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United States Environmental Protection Agency
Office of Water
Office of Environmental Information
Washington, DC
EPA-843-R10-002
National Wetland Condition Assessment
Laboratory Operations
Manual
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National Wetlands Condition Assessment
Laboratory Operations Manual Page
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National Wetlands Condition Assessment
Laboratory Operations Manual Page ill
TABLE OF CONTENTS
1. INTRODUCTION 1
1.1 Form Logistics 2
1.2 Tracking Samples 2
1.3 Sending Resultant Data Forms 3
1.4 References 3
2. WATER CHEMISTRY 4
2.1 Introduction to Indicator 4
2.2 Parameters for the NWCA 4
2.3 Performance-based Methods 5
2.4 Sample Processing and Preservation 5
2.5 Quality Assurance/Quality Control (QA/QC) Procedures 7
2.5.1 Internal QA/QC Procedures 8
2.6 References 8
3. ALGAE 9
3.1 I ntroduction to I ndicator 9
3.1.1 References 11
3.2 Subsampling 11
3.2.1 Safety Precautions 11
3.2.2 Recommended Apparatus/Equipment 12
3.2.3 Methods 13
3.2.4 Quality Assurance/Quality Control 13
3.2.5 References 14
3.3 Diatom Cleaning 14
3.3.1 Definitions 14
3.3.2 Safety Precautions 15
3.3.3 Recommended Apparatus/Equipment 15
3.3.4 Methods 15
3.3.5 Quality Assurance/ Quality Control 17
3.3.6 References 17
3.4 Preparation of Diatom Slides using Naphrax™ Mounting Medium 17
3.4.1 Safety Precautions 18
3.4.2 Recommended Apparatus/ Equipment 18
3.4.3 Methods 19
3.4.4 Quality Assurance/ Quality Control 23
3.4.5 References 23
3.5 Analysis of Diatoms 24
3.5.1 Recommended Apparatus/ Equipment 24
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3.5.2 Methods 24
3.5.3 Quality Assurance/Quality Control 26
3.5.4 References 26
3.6 Analysis of All Algae 28
3.6.1 Definitions 28
3.6.2 Recommended Apparatus/ Equipment 29
3.6.3 Methods 29
3.6.4 Calculations and Reporting 32
3.6.5 Quality Assurance/ Quality Control 33
3.6.6 References 33
3.7 Phytoplankton Chlorophyll a 33
3.7.1 Summary of Method 35
3.7.2 Definitions 35
3.7.3 I nterferences 35
3.7.4 Safety Precautions 36
3.7.5 Equipment and Supplies 36
3.7.6 Reagents and Standards 37
3.7.7 Sample Collection, Preservation and Storage 38
3.7.8 Calibration and Standardization 39
3.7.9 Procedure 40
3.7.10 Data Analysis and Calculations 41
3.7.11 Method Performance 42
3.7.12 Pollution Prevention 42
3.7.13 Waste Management 42
3.7.14 Quality Control 43
3.7.15 References 43
3.8 Microtiter Plate Enzyme-Linked Immuno-Sorbent Assay (ALISA) for Microcystin 44
3.8.1 Definitions 44
3.8.2 Safety Precautions 45
3.8.3 Equipment and Supplies 45
3.8.4 Procedures 46
3.8.5 Quality Control 51
3.8.6 References 52
3.9 Taxonomic Standards Protocols 52
3.9.1 Images 54
3.9.2 Table of References 55
3.9.3 Taxonomic Harmonization 55
3.9.4 Final Products 56
3.10 Pertinent QA/QC Procedures 57
4. VEGETATION 58
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4.1 Introduction to Indicator 58
4.2 Receiving Voucher Samples 58
4.2.1 Definitions 58
4.2.2 Tracking information 60
4.3 Supplies and Equipment for Sample Handling 63
4.4 Handling Vegetation Samples 63
4.4.1 Plant Sample ID Tag and Label Form 63
4.4.2 Drying Samples 65
4.4.3 Treat samples for detritivores, molds, and pests 66
4.5 Identification of Vegetation Samples 66
4.5.1 Taxonomic standard 66
4.5.2 Species Identifications for Voucher Samples 67
4.6 Mounting and Storing Herbarium Sheets 67
4.7 Quality Assurance 67
4.8 References 68
5. SOIL 70
5.1 I ntroduction to Indicator 70
5.2 Quality Assurance/Quality Control 70
5.3 Receiving Regulated Soils 71
5.4 Parameters for the NWCA 74
5.5 Laboratory Sample Collection and Preparation 78
5.5.1 Summary of Method 79
5.5.2 I nterferences 79
5.5.3 Safety 80
5.5.4 Equipment 80
5.5.5 Reagents 81
5.5.6 Procedures 81
None - No visual efferevescence 82
5.5.7 Calculations 84
5.5.8 Report 84
5.5.9 Precision and Accuracy 84
5.5.10 References 85
5.6 Sampling Archiving 85
5.7 PSDA Air-dry, <2mm Particles 86
5.7.1 Summary of Method 86
5.7.2 Interferences 86
5.7.3 Precautions 87
5.7.4 Equipment/Materials 87
5.7.5 Reagents 89
5.7.6 Procedure 89
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5.7.7 Calculation Error! Bookmark not defined.
5.7.8 Reporting 92
5.7.9 References 92
5.8 PSDA Pipet Analysis, Particles <2 mm 94
5.8.1 Summary of Method 94
5.8.2 Interferences 96
5.8.3 Safety 98
5.8.4 Equipment 98
5.8.5 Reagents 99
5.8.6 Procedures 100
5.8.7 Calculations 106
5.8.8 Report 108
5.8.9 Precision and Accuracy 108
5.8.10 References 108
5.9 Water Retention, 1500-kPa 110
5.9.1 Summary of Method 110
5.9.2 Interferences 110
5.9.3 Safety 111
5.9.4 Equipment 111
5.9.5 Reagents 111
5.9.6 Procedure 111
5.9.7 Calculations 112
5.9.8 Report 113
5.9.9 Precision and Accuracy 113
5.9.10 References 113
5.10 Calcium Carbonate Equivalent 114
5.10.1 Summary of Method 114
5.10.2 Interferences 114
5.10.3 Precautions 114
5.10.4 Equipment/Materials 115
5.10.5 Reagents 115
5.10.6 Procedure 115
5.10.7 Calculation 116
5.10.8 Reporting 117
5.10.9 References 117
5.11 Total Carbon, Total Nitrogen, and Total Sulfur 118
5.11.1 Summary of Method 118
5.11.2 Interferences 119
5.11.3 Precautions 119
5.11.4 Equipment/Materials 119
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5.11.5 Reagents 120
5.11.6 Procedure 120
5.11.7 Calculation 121
5.11.8 Reporting 122
5.11.9 References 122
5.12 pH 123
5.12.1 Summary of Method 123
5.12.2 Interferences 124
5.12.3 Precautions 124
5.12.4 Equipment/Materials 124
5.12.5 Reagents 124
5.12.6 Procedure 125
5.12.7 Calculation 125
5.12.8 Reporting 126
5.12.9 References 126
5.13 BaCI2-Triethanolamine, pH 8.2 Extraction 126
5.13.1 Summary of Method 126
5.13.2 Interferences 126
5.13.3 Safety 126
5.13.4 Equipment 127
5.13.5 Reagents 127
5.13.6 Procedure 128
5.13.7 Calculations 129
5.13.8 Report 129
5.13.9 Precision and Accuracy 129
5.13.10 References 130
5.14 Cation Exchange Capacity (Total) 130
5.14.1 Summary of Method 130
5.14.2 Interferences 130
5.14.3 Safety 130
5.14.4 Equipment 131
5.14.5 Reagents 131
5.14.6 Procedure 132
5.14.7 Calculations 134
5.14.8 Report 135
5.14.9 References 135
5.15 Cation Exchange Capacity (Ca, Mg, K, Na) 135
5.15.1 Summary of Method 135
5.15.2 Interferences 136
5.15.3 Precautions 136
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5.15.4 Equipment/Materials 136
5.15.5 Reagents 136
5.15.6 Procedure 138
5.15.7 Calculation 139
5.15.8 Reporting 140
5.15.9 References 140
5.16 Acid Oxalate Extraction (Al, Fe, Mn, P, Si) 140
5.16.1 Summary of Method 140
5.16.2 I nterferences 140
5.16.3 Precautions 141
5.16.4 Equipment/Materials 141
5.16.5 Reagents 141
5.16.6 Procedure 143
5.16.7 Calculation 146
5.16.8 Reporting 147
5.16.9 References 147
5.17 Electrical Conductivity (Saline Prediction) 148
5.17.1 Summary of Method 148
5.17.2 Interferences 148
5.17.3 Precautions 148
5.17.4 Equipment/Materials 148
5.17.5 Reagents 148
5.17.6 Procedure 149
5.17.7 Calculation 149
5.17.8 Reporting 149
5.17.9 References 150
5.18 Citrate Dithionite Extraction (Al, Fe, and Mn) 150
5.18.1 Summary of Method 150
5.18.2 Interferences 150
5.18.3 Precautions 150
5.18.4 Equipment/Materials 151
5.18.5 Reagents 151
5.18.6 Procedure 153
5.18.7 Calculation 155
5.18.8 Reporting 155
5.18.9 References 156
5.19 Phosphorus (Olsen) 156
5.19.1 Summary of Method 156
5.19.2 Interferences 156
5.19.3 Precautions 156
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5.19.4 Equipment/Materials 157
5.19.5 Reagents 157
5.19.6 Procedure 158
5.19.7 Calculation 159
5.19.8 Reporting 160
5.19.9 References 160
5.20 Phosphorus (Mehlich No. 3) 160
5.20.1 Summary of Method 160
5.20.2 Interferences 160
5.20.3 Precautions 161
5.20.4 Equipment/Materials 161
5.20.5 Reagents 161
5.20.6 Procedure 162
5.20.7 Calculation 163
5.20.8 Reporting 164
5.20.9 References 164
5.21 Trace Elements (Ag, Ba, Be, Cd, Co, Cr, Cu, Hg, Mn, Mo, Ni, P, Pb, Sb, Se, Sn, Sr,
Tl, V, W, andZn) 164
5.21.1 Summary of Method 164
5.21.2 Interferences 165
5.21.3 Precautions 165
5.21.4 Equipment/Materials 165
5.21.5 Reagents 166
5.21.6 Procedure 166
5.21.7 Calculation 173
5.21.8 Reporting 174
5.21.9 References 174
5.22 Bulk Density (Hammered Core, Field State) 175
5.22.1 Summary of Method 175
5.22.2 Interferences 176
5.22.3 Equipment 176
5.22.4 Procedure 176
5.22.5 Calculations 177
5.22.6 Report 177
5.22.7 References 177
5.23 Bulk Density (Saran Clod, Field State) 177
5.23.1 Summary of Method 178
5.23.2 Interferences 178
5.23.3 Precautions 179
5.23.4 Equipment 179
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5.23.5 Reagents 180
5.23.6 Procedure 180
5.23.7 Calculation 181
5.23.8 Reporting 182
5.23.9 References 182
5.24 Bulk Density (Oven-Dry, clods and cores) 183
5.24.1 Summary of Method 183
5.24.2 Interferences 183
5.24.3 Safety 184
5.24.4 Equipment 184
5.24.5 Reagents 185
5.24.6 Procedure 185
5.24.7 Calculations 186
5.24.8 Report 187
5.24.9 References 187
5.25 Bulk Density (33-kPA Desorption, clods only) 188
5.25.1 Summary of Method 188
5.25.2 Interferences 188
5.25.3 Safety 189
5.25.4 Equipment 189
5.25.5 Reagents 190
5.25.6 Procedure 190
5.25.7 Calculations 192
5.25.8 Report 193
5.25.9 References 193
Appendix A 195
Appendix B 203
Appendix C 243
Appendix D 254
Pressing Samples 255
Mounting Plant Samples 257
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Laboratory Methods Manual Page 1 of 256
1. INTRODUCTION
This manual describes laboratory protocols and is for use by laboratory personnel for samples
associated with the National Wetland Condition Assessment (NWCA). The NWCA is one in a
series of National Aquatic Resource Surveys (NARS) conducted by the Environmental
Protection Agency (USEPA) to provide the public with a comprehensive assessment of the
condition of the Nation's waters. In addition to wetlands, the NARS will assess coastal waters,
lakes, rivers, and streams in a revolving sequence.
USEPA will collaborate with state, tribal, federal, and other partners to implement the NWCA to
meet three goals:
1. Produce a report that describes the ecological condition of the Nation's wetlands.
2. Assist states and tribes in the implementation of wetland monitoring and assessment
programs that will guide policy development and aid decision-making.
3. Advance the science of wetlands monitoring and assessment to support management
needs.
The NWCA responds to the long-term goals outlined in USEPA's current strategic plan (USEPA
2006a) to improve the Nation's water quality (Goal 2.3) and to protect, sustain, and restore the
health of critical natural habitats and ecosystems, including wetlands (Goal 4.3). Development
of the NWCA will build on the accomplishments of the U.S. Fish and Wildlife Service (USFWS)
and their production of national reports on status and trends in wetland acreage. When taken
together, the NWCA and the USFWS Wetland Status and Trends (S&T) results will be used to
measure progress toward attainment of the national goal to increase the quantity and quality of
the Nation's wetlands. These complementary studies can influence how wetlands are managed
at local, state, and national scales (Scozzafava, et. al. 2007).
The complete documentation of overall project management, design, methods, and standards
for the NWCA is contained in four companion documents:
1. National Wetland Condition Assessment: Quality Assurance Project Plan (EPA-843-R10-
003)
2. National Wetland Condition Assessment: Site Evaluation Guidelines (EPA-843-R10-004)
3. National Wetland Condition Assessment: Field Operations Manual (EPA-843-R10-001)
4. National Wetland Condition Assessment: Laboratory Operations Manual (EPA-843-R10-
002)
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This document (Laboratory Operations Manual) contains information on the methods for
analyses of the samples collected during the project, quality assurance objectives, sample
handling, and data reporting. Methods described in this document are to be used specifically in
work relating to NWCA. All Project Cooperator laboratories should follow these guidelines.
Mention of trade names or commercial products in this document does not constitute
endorsement or recommendation for use. More details on specific methods for site evaluation,
sampling, and sample processing can be found in the appropriate companion document.
The suggested citation for this document is:
USEPA. 2011. National Wetland Condition Assessment: Laboratory Operations Manual. EPA-
843-R10-002. U.S. Environmental Protection Agency, Office of Water and Office of
Research and Development, Washington, DC.
1.1 Form Logistics
The National Wetland Condition Assessment will include laboratory processing of some
indicators. Water chemistry, algae, vegetation, and soils samples will be sent to laboratories for
analysis. These laboratories should fill out the applicable data forms (appendices) and email
them to the Information Management Coordinator at NARSDataSubmission@epa.gov. If email
is not available, forms may be shipped to:
Marlys Cappaert, SRA Intl.
c/o EPA
200 SW 35th Street
Covallis, OR 97331
1.2 Tracking Samples
With the samples from the field crews, laboratories should also receive tracking forms (refer to
NWCA Field Operations Manual (FOM)). These forms will list the samples/specimens that
should be included in the shipment. Laboratory personnel should cross check the forms with
the samples received to verify that there are not any inconsistencies. If any sample/specimen is
missing from the shipment, contact the Information Management Coordinator (541-754-4663)
immediately.
The lab will adhere to strict sample tracking procedures to ensure samples are handled in an
expeditious manner. Tracking procedures include, at a minimum, immediate notification of
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shipment arrival followed within 24 hours by an electronic listing of the samples received.
Results are reported in an electronic format detailed in the MAIA-Estuaries Data Format
Manual. A copy of the cover letter which accompanies the results shall be sent to the EPA
Contracting Officer.
1.3 Sending Resultant Data Forms
Resultant data files (i.e., EXCEL spreadsheets) are also emailed to the Information
Management Coordinator at NARSDataSubmission(S)epa.gov for the purpose of quality
assurance. A written report detailing the methods used discussing problems encountered must
be included as an appendix to this report. Original records, such as laboratory notebooks, shall
be retained for at least five years following final report submission, as they may be requested to
be sent to EPA. If email is not available, forms the report may be shipped to:
Marlys Cappaert, SRA Intl.
c/o EPA
200 SW 35th Street
Covallis, OR 97331
1.4 References
USEPA. (1997) MAIA-Estuaries 1997 Laboratory Data Format and Transmittal Guidance
Document. United States Environmental Protection Agency, Atlantic Ecology Division.
Narragansett, Rl.
Scozzafava, M.E., I.E., Dahl, C., Faulkner, and M. Price. 2007. Assessing status, trends, and
condition of wetlands in the United States. National Wetlands Newsletter 29:24-28.
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2. WATER CHEMISTRY
2.1
Introduction to Indicator
Along with vegetation and soil, water is one of the critical media for defining a wetland. Some
studies show that water chemistry analyses are useful for evaluating wetland ecological integrity
and evaluating stressor-response relationships (Lane and Brown, 2007; Reiss and Brown,
2005). Categories of data to be collected include the following:
1. pH
2. Nutrient Enrichment
3. Dissolved oxygen
4. Temperature
For more detailed information please see the report Ecological Indicators for the 2011 National
Wetland Condition Assessment (in preparation).
2.2
Parameters for the NWCA
A total of four parameters will be measured from each water sample collected (Table 2.2-1).
Table 1 also lists example methods that have been demonstrated to achieve the required
objectives. Figure 1 provides a flow diagram of water chemistry processing procedures.
Table 2.2-1. Water chemistry parameters measured for the National Wetland Condition
Assessment.
Analyte
Conductivity
PH
Ammonia (NH3)
Units
uS/cm at
25°C
pH units
mgN/L
Example Method(s)1
EPA 120.6
(conductivity cell)
EPA 150. 6 (modified)
(Aliquot from bulk water sample)
EPA 350.1 , or modifications
Method Detection
Limit Objective2
NA
NA
0.02
(e.g., Automated Colorimetric, or modified to
use salicylate and dichloroisocyanurate with
analysis by flow injection analyzer (FIA))
Nitrate-Nitrite (NO3~ mg N/L
N02-)
EPA 353.2 (modified)
(Automated colorimetric analysis with cadmium
reduction by FIA)
0.02
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Table 2.2-1. Water chemistry parameters measured for the National Wetland Condition
Assessment.
Analyte
Total Nitrogen (TN)
Units
mg/L
Example Method(s)1
EPA 353.2 (modified)
(Persulfate Digestion; Automated colorimetric
analysis with cadmium reduction; modified for
analysis by FIA)
Method Detection
Limit Objective2
0.01
Total Phosphorous ug P/L EPA 365.1 (modified) Persulfate digestion 2
(TP)
1 Methods presented are those used for WSA analyses (and have met the performance characteristics listed). In some cases, a
potential alternative method is also presented. Methods presented here should not be interpreted as the required method(s) to
be used by all laboratories analyzing NWCA samples.
2 The method detection limit is determined as a one-sided 99% confidence interval from repeated measurements of a low-level
standard across several calibration curves.
2.3 Performance-based Methods
As an alternative to specifying laboratory methods for sample analysis, we use a performance-
based approach that defines a set of laboratory method performance requirements for data
quality. Method performance requirements for this project identify detection limit, precision, and
accuracy objectives for each parameter (Table 2.3-1). It should be noted that there are no
required methods to be used by all participating laboratories. Laboratories may choose to use
other analytical methods for any target analytes as long as they are able to achieve the same
performance requirements as listed in Table 2.3-1.
2.4 Sample Processing and Preservation
Upon receipt of samples, inspect each sample and complete the tracking form before storing in
the refrigerator at 4°C until aliquots are ready to be prepared. Figure 1-1 illustrates the sample
preparation procedures, including filtering and acidifying, for the various analytes.
Filter the aliquots for ammonia through 0.4um pore size polycarbonate filters within 48 hours of
arrival at the Laboratory. Rinse vacuum filter funnel units thoroughly with reverse-osmosis (RO)
or de-ionized (Dl) water five times before each use and in between samples. After placing a
filter in the funnel unit, run approximately 100 ml of RO or Dl water through the filter, with
vacuum pressure, to rinse the filter. Discard the rinse water, then place the appropriate sample
bottle under the funnel unit and filter the sample directly into the bottle. If a new filter is needed,
remove the sample bottle, and rinse the new filter with 100 ml of RO or Dl water before
continuing sample filtration.
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After all filtered and unfiltered aliquots are collected add ultra-pure acid (H2SO4) to the sample in
the aliquot container. Store all aliquots in a refrigerator at 4°C for a holding time of 28 days.
Table 2.3-1 : Performance requirements for water chemistry analytical methods
Analyte Units
Conductivity |^S/cm at
25° C
pH pH units
Ammonia (NH3) mg N/L
Nitrate-Nitrite mg N/L
(N03-N02)
Total Nitrogen mg/L
(TN)
Total |jg P/L
Phosphorus (TP)
Potential
Range
of Samples1
1 to 15,000
3.7 to 10
Oto 17
0 to 360 (as
nitrate)
0.1 to 90
0 to 22,000
Long -Term
MDL
Objective2
NA
NA
0.01
(0.7 ueq/L)
0.01
0.01
2
Laboratory
Reporting Limit3
2.0
NA
0.02
(1.4 ueq/L)
0.02
0.02
4
Transition
Value4
20
5.75
and>8.25
0.10
0.10
0.10
20
Precision
Objective5
±2 or ±10%
± 0.08 or ±
0.15
± 0.01 or
±10%
± 0.01 or
±10%
± 0.01 or
±10%
±2 or ±10%
Bias
Objective6
± 2 or 5%
±0.05 or±
0.10
±0.01 or
±10%
±0.01 or
±10%
±0.01 or
±10%
±2 or ±10%
1 Estimated from samples analyzed at the WED-Corvallis laboratory between 1999 and 2005 for TIME, EMAP-West, and WSA
streams from across the U.S.
2 The long-term method detection limit is determined as a one-sided 99% confidence interval from repeated measurements of a
low-level standard across several calibration curves, based on USGS Open File Report 99-193. These represent values that
should be achievable by multiple labs analyzing samples over extended periods with comparable (but not necessarily identical)
methods.
3 The minimum reporting limit is the lowest value that needs to be quantified (as opposed to just detected), and represents the
value of the lowest nonzero calibration standard used. It is set to 2xthe long-term detection limit, following USGS Open File
Report 99-193 New Reporting Procedures Based on Long-Term Method Detection Levels and Some Considerations for
Interpretations of Water-Quality Data Provided by the U.S. Geological Survey National Water Quality Laboratory.
4 Value at which performance objectives for precision and bias switch from absolute (< transition value) to relative 9> transition
value). Two-tiered approach based on Hunt, D.T.E. and A.L. Wilson. 1986. The Chemical Analysis of Water: General Principles
and Techniques. 2nd ed. Royal Society of Chemistry, London, England.
5 For duplicate samples, precision is estimated as the pooled standard deviation (calculated as the root-mean square) of all
samples at the lower concentration range, and as the pooled percent relative standard deviation of all samples at the higher
concentration range. For standard samples, precision is estimated as the standard deviation of repeated measurements across
batches at the lower concentration range, and as percent relative standard deviation of repeated measurements across batches
at the higher concentration range.
6 Bias (systematic error) is estimated as the difference between the mean measured value and the target value of a performance
evaluation and/or internal reference samples at the lower concentration range measured across sample batches, and as the
percent difference at the higher concentration range.
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Sample Receipt
4-L Bulk Samples
•Inspect samples anocomplete tracking form
•Store at 4°C in darkness
J
Filtration
1
Process within 48 Hours
(0.4 um)
i • i
•HOPE bottle
•Not acid
washed
•No preservative
y
i
Analyses:
•Nitrate
•Nitrite
(7 day holding
time)
r
1
1
\
mi
•HOPE bottle
•Acid washed
•Preserve with
H2S04
V
•
^
-
Analyses:
•Ammonia
(14 day
holding time)
1
r -
1
Not Filtered
< ^
•HOPE bottle
•Acid washed
•Preserve with
H2S04
V
J
fc
•HOPE bottle
•Not acid washed
•No preservative
J ^
*
Analyses:
•Total Phosphorous
•Total Nitrogen
(28 day holding
time)
^
Analyses:
•Conductivity
(7 day holding
time)
•pH
(3 day holding
timel
Figure 1-1. Water chemistry sample processing procedures.
2.5 Quality Assurance/Quality Control (QA/QC) Procedures
Since multiple laboratories will be analyzing water chemistry samples, specific quality control
procedures must be implemented to ensure that 1) the data quality objectives are being met and
2) data is consistent and comparable among all participating labs. Specific QA/QC procedures
will depend on whether or not the lab has been certified by the National Environmental
Laboratory Accreditation Conference (NELAC). More stringent QA/QC requirements will be
implemented for labs that are not NELAC certified.
A total of 10% of the samples collected will be re-analyzed for quality control. Re-analysis will
be performed by another experienced technician at an independent laboratory who did not
participate in the original analyses. EPA estimates that approximately 100 samples may
require re-analysis for all analytes. EPA will inform the laboratories which random samples will
be re-analyzed. The QC laboratory technician should complete another copy of the Water
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Laboratory Methods Manual Page 8 of 256
Chemistry Datasheet for each sample. Each sheet should be labeled with the term "QC Re-
analysis." As each sheet is completed, it should be emailed to the Information Management
Coordinator at NARSDataSubmission@epa.gov.
EPA will compare the results generated by the primary and QC laboratories for each sample
and calculate percent similarity (refer to Chapter 2 of the QAPP). It is expected that the two
analyses should have a similarity of >90%. If not, the reasons for the discrepancies between
laboratories should be discussed. Results less than these values will be investigated and
logged for indication of error patterns or trends.
Significant differences may result in the re-analysis of samples by the primary laboratory and a
second QC check by the secondary laboratory. All samples must be stored at the laboratory
until the Information Management Coordinator notifies the lab.
2.5.1 Internal QA/QC Procedures
Prior to sample analysis, all laboratories will be required to provide internal QA documentation
(e.g., Quality Management Plan, Quality Assurance Project Plan) for external review. QA
documentation will be reviewed by the QA Coordinator to ensure consistency among all
participating labs.
2.6 References
Hunt, D.T.E. and A.L. Wilson. 1986. The Chemical Analysis of Water: General Principles and
Techniques. 2nd ed.. Royal Society of Chemistry, London, England.
Lane, C.R. and M.T. Brown. 2007. Diatoms as indicators of isolated herbaceous wetland
condition in Florida, USA. Ecological Indicators 7(3):521-540.
Reiss, K.C. and M.T. Brown. 2005. The Florida Wetland Condition Index (FWCI):
developing biological indicators for isolated depressional forested wetlands. A Report
to the Florida Department of Environmental Protection. Howard T. Odum Center for
Wetlands, University of Florida, Gainesville, Florida, USA.
Youden, W.J. 1969. Ranking laboratories by round-robin tests. In Precision Measurement and
Calibration. H.H. Ku, ed. NBS Special Publication 300, Vol. 1. U.S. GPO Washington,
D.C.
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3. ALGAE
3.1 Introduction to Indicator
Algae, which include planktonic (open water), benthic (periphyton), and metaphyton forms, are
an extremely important ecosystem component providing essential primary productivity as well
as a resource for higher level organisms including macroinvertebrates and fish (Mitsch and
Gosselink, 2007). Like other biotic taxa, many indicator attributes or metrics can be derived
from data describing taxonomic composition and abundance of algae. Categories of data to be
collected include the following:
1. Species composition and abundance
2. Guilds
3. Productivity
4. Habitat evaluation
5. Physiology
For more detailed information please see the report Ecological Indicators for the 2011 National
Wetland Condition Assessment (in preparation).
Algae samples will be collected by field crews and sent to a laboratory for analysis on a regular
basis during the field season to avoid delays in processing and identifying samples (for more
information on shipping field samples, refer to FOM section 12.3). Laboratories should receive
a 250ml composite sample of algae from benthic and vegetative surfaces that includes both
diatoms and soft algae. This sample will be subsampled so the diatoms can be acid cleaned
and analyzed separately from the subsample of composite algae that includes soft algae along
with the diatoms (see Figure 3.1-1 for a flowchart of processing activities). Crews will also send
a phytoplankton chlorophyll a sample from those assessment areas where standing water is
present. This sample will have been filtered using a 0.4um pore size glass filter in the field.
Laboratories should receive the filter in a small vial that has been frozen. The following method
for the analysis of composite algae samples has been adapted from Protocols for the analysis of
algal samples collected as part of the U. S. Geological Survey National Water-Quality
Assessment program (Charles et al., 2002) and is used to identify and enumerate algal
populations. EPA recommends these procedures for the analysis of NWCA composite algae
samples. However, we recognize that in some cases there are alternate methods that may
produce comparable results. Laboratories interested in using alternate methods must submit
them to EPA for approval prior to conducting any analyses (see QAPP for lab and personnel
qualifications).
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Chlorophyll a
sample
and filter
Centrifuge
sample
Freeze
sampleat-
20*C
Extract
pigment
Centrifuge
sample
Prepare
aliquots
Measure
fluorescence
Sample Receipt:
•250 ml composite algal
sample
•Chlorophyll a sample and
filter
250 ml composite
algal sample
X ml composite
subsample
X ml diatom
subsample
Acid clean
diatoms
Palmer-Maloney
Prep, (all algae)
Prepare diatom
mounts
Analyze
composite algae
Analyze
diatoms
Figure 3.1-1: Soft algae, diatom, and Chlorophyll-a processing procedure.
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3.1.1 References
Charles, D. F., C. Knowles, and R. Davis. 2002. Protocols for the analysis of algal samples
collected as part of the U.S. Geological Survey National Water-Quality Assessment
Program. Patrick Center for Environmental Research Report No. 02-06. Patrick Center
for Environmental Research, The Academy of Natural Sciences, Philadelphia,
Pennsylvania.
Mitsch, W.J.,and J.G.Gosselink. 2007. Wetlands. 4th ed. New York:Wiley.
Stevenson, Jan. Personal communication August 11, 2009.
USEPA. 2008. National Rivers and Streams Assessment: Laboratory Methods Manual. EPA
841-B-07-010. U.S. Environmental Protection Agency, Office of Water and Office of
Research and Development, Washington, DC.
USEPA. 2006. Survey of the Nation's Lakes. Laboratory Methods Manual. EPA841-B-06-005.
U.S. Environmental Protection Agency, Washington, DC.
3.2 Subsampling
Qualitative composite algal samples must be subsampled carefully to ensure that all algal forms
are represented in the subsamples to be analyzed. The sample received from field crews may
have large amounts of sand, silt, or other heavy material that can interfere with algal analysis.
Therefore, this is a special procedure involving the subsampling of these samples. In this
procedure, the liquid portion is subsampled by volume and the heavier material that is difficult to
suspend is separated by mass. Protocols for subsampling samples with heavy sediments can
be found in the NAQWA protocols (section 7.5, pages 20-21).
3.2.1 Safety Precautions
Samples will arrive at the lab in a Lugol's preservative (see Algae Chapter of FOM). Samples
preserved with the appropriate amount of Lugol's solution will look like weak tea (Figure 3.2-1).
If a sample does not contain a preservative call the Information Management Coordinator (541-
754-4663) before handling. Upon receipt at the lab, add a few (1-3) drops of 100% buffered
formalin to the samples to ensure preservation.
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Figure 3.2-1: Image of algal samples preserved with Lugol's. The middle vial contains the appropriate
amount of Lugol's while the left contains too little and the right contains too much. (Photograph from
LaLiberte, 2010)
3.2.2 Recommended Apparatus/Equipment
• Distilled (DW) or reverse osmosis (RO) water
• Dispenser bottle for DW or RO water
• Beakers
• Beaker holding box (24 slots)
• Graduated cylinders (10 ml, 25 ml, 100 ml, 250 ml, 500 ml and 1 L)
• 20-ml vials
• 20-ml vial with 1 ml intervals marked on it for measuring volumes
• Diamond scriber
• Protective clothing (gloves, lab coat or apron, eye protection)
• Positive-draw fume hood
• Spatulas
• Plastic disposable pipettes (tips of plastic pipettes can be cut)
• Small turkey baster
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3.2.3 Methods
1. The 250ml composite sample received from field crews must be separated into 2
subsamples for separate counting of diatoms and of all algae. Deciding how much to
subsample for all algae and diatoms is somewhat of a judgment call. If the sample is very
silty, it will be better to subsample a minimum of 10 ml for all algae, whereas if the sample
looks clear, at least 20 ml is recommended. Sample at least 10 ml for soft and at least 20 ml
for diatoms.
2. If there are visible macroalgal forms, these should have been cut into smaller pieces by the
field crews and included in proper proportions in aliquots. If macroalgae were not cut by
field crews, then they should be cut before subsampling.
3. Suspend the algal material by shaking or swirling and carefully pour the determined amount
of subsample into a graduated cylinder. Ensure the sample stays well mixed during
subsampling. Alternatively, use a wide-bore pipette or turkey baster to subsample the
125ml composite sample, which allows pumping the pipette or baster to mix the sample,
while withdrawing an aliquot. Select a method that allows a representative subsample.
Filaments and debris will block pipettes with small openings. Careful pouring or turkey
basters may be more appropriate. Withdrawing multiple aliquots for each sample, each
approximately 5 ml, enables drawing 2-4 aliquots for each 10-20 ml diatom and all-algae
subsample.
4. Transfer each subsample to the 20-ml vial labeled with Subsample ID#.
5. Record the subsample volume and beaker ID of each subsample on Subsample Data
Sheet.
3.2.4 Quality Assurance/Quality Control
Since algae are microscopic, there is significant possibility of contamination of samples.
Laboratory rooms where raw samples are subsampled should be kept as clean as possible. Lab
bench surfaces should be kept clean and free of debris.
The appropriate size of graduated cylinder for measuring samples and subsamples is critical.
The sample should be at least one-third the capacity of graduated cylinder and the units of the
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graduated cylinder should allow estimation to the nearest milliliter (finer for the small graduated
cylinders used to measure a portion of a subsample).
3.2.5 References
Moulton, S.R., II, J.G. Kennen, R.M. Goldstein, J.A. Hambrook. 2002. Revised protocols for
sampling algal, invertebrate, and fish communities in the National Water-Quality
Assessment program, U.S. Geological Survey Open-File Report 02-150. In Press.
Porter, S.D., T.F. Cuffney, M.E. Gurtz, M.R. Meador. 1993. Methods for collecting algal samples
as part of the National Water-Quality Assessment program. U.S. Geological Survey
Open-File Report 93-409, Raleigh, NC [39 pp] http://water.usgs.gov/nawqa/
protocols/OFR-93-409/alg1 .html
United States Geological Survey, National Water-Quality Assessment Program. 1997.
Procedures for Processing NAWQA Algal Samples. Draft Manuscript. February 1997.
3.3 Diatom Cleaning
This protocol will apply to the subsample which will be analyzed for diatoms only. To identify
and enumerate diatoms accurately at the species and variety levels, it is necessary to remove
both extracellular and intracellular organic matter from the siliceous frustules of diatoms and
other material such as the soft algae in the sample. Removing the organic matter is necessary
so that all details of diatom structures essential to taxonomic identification are clearly visible.
This protocol describes a method for removing organic material from a sample by digesting it
with nitric acid.
3.3.1 Definitions
Digestion in this procedure refers to the solubilization of organic material by strong acid
oxidation.
Diatom cells, called frustules, are composed of two valves. They have a siliceous structure,
the features of which are used for taxonomic identification.
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3.3.2 Safety Precautions
Nitric acid is an extremely hazardous reagent. As a strong acid oxidizer it can cause severe
burning of exposed skin and clothing. At room temperature, concentrated nitric acid produces
intense fumes when exposed to open air. Any concentrated nitric acid containers open to the
air must be contained within a positive-draw fume hood at all times. There are no exceptions to
this rule. Personnel are required to wear safety glasses, protective gloves and lab coats at all
times when handling concentrated nitric acid. This is especially important when handling/venting
the digestion vessels.
NWCA Samples delivered to the Diatom Preparation Laboratory will have been prepared with
Lugol's solution (see section 3.2.2 of LOM Algae Chapter of FOM). This information should be
on the shipping forms included with the samples or affixed to the shipping container.
Hot plate temperatures required for this procedure are high enough to cause severe burning of
exposed skin. Use extreme care when manipulating slides on the hot plate and when working
close to the hot plate.
3.3.3 Recommended Apparatus/Equipment
• Hot plate
• Positive-draw fume hood
• Safety glasses
• Acid-impervious hand protection
• Laboratory coat or apron, acid resistant
• Dl water
• Siphon
3.3.4 Methods
1. The diatom subsample is digested under the hood on a hot plate.
a. As an alternative the subsample can be digested in test tubes and heated in a
sand bath.
b. As another alternative the ANSP microwave digestion method can also be used
(protocol P-13-42).
2. Set the hot plate on 250°C.
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3. Cautiously add at least 50 ml nitric acid (under the hood) to the sample (rule minimum: 1:2
sample to acid ratio).
a. Hydrogen Peroxide may be used as a less toxic alternative to nitric acid.
4. Heat the acid and sample mixture. Boil for 1 to 2 hours. Add Dl water as needed to maintain
the liquid levels.
5. Move the sample from the hot plate to a tray and let it cool for a few hours.
6. Very carefully add Dl water (under the hood) up to the top of the beaker. Leave this
overnight in the hood (for at least 8 hours).
a. As an alternative, centrifuge samples for 10 minutes at approximately 2000 rpm.
7. Transfer the tray to the "water changing station", siphon the water from the center of the
water column (from side to side and top to bottom) to around 100 ml (the mark to which you
remove water at this point can be variable). Avoid siphoning diatoms settled on the bottom
or adsorbed to the sides of the beaker and water surface.
Note: Do not disturb the settled layer of sample on the bottom so none of the sample is lost.
Also, diatoms adsorb to sides of the beaker and water surface at the top, so after siphoning add
Dl water, washing the sides of the beaker with a Dl water bottle. Sprinkle water on the surface
to dislodge algae adsorbed to the water surface. Leave the sample for at least 8 hours to allow
time for the diatoms to settle in the beaker before washing again (the rule is 1 hour per 1
centimeter so the smallest diatoms can settle).
8. After a neutral pH is achieved (approximately 6-8 washings if samples are settled, 4-5 if
samples are centrifuged), the digested diatom material is transferred to the pre-labeled vial
for cleaned diatom material.
Note: The volume in the "diatom volume after digestion" vial is a judgment call by the laboratory
technician. The goal is to achieve an even distribution for the slides in the next step. Tip: when
shaken, the vial needs to be transparently cloudy. Leave some space for dilution in that vial. If
the sample looks completely clear after digestion assume it is a sparse sample, and try to very
carefully concentrate the whole sample to about 5 ml. It is easier to play with the volume than to
count samples that are too sparse.
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3.3.5 Quality Assurance/ Quality Control
Diatom frustules are microscopic, generally falling in the fine silt size range; therefore, there is a
possibility that samples can be contaminated. Laboratory rooms where raw or processed
samples are handled should be kept as clean as possible. Lab bench surfaces should be kept
clean and free of debris. Techniques similar to those used for sterile experiments
(bacteriological plating, etc.) should be followed to minimize the risk of cross contamination of
samples. Where feasible, disposable pipettes, stirrers, etc. should be used. Where they cannot,
they should be rinsed in DW or RO water at least three times after each sample. (Explanatory
note: at times, tap water, because of algal blooms and use of diatomaceous earth filters, may
contain diatoms.) All equipment should be stored dry to prevent growth of algae or fungi.
3.3.6 References
Acker, F., B. Russell, and E. Hagan. 2002. Protocols for the analysis of algal samples collected
as part of the U.S. Geological Survey National Water-Quality Assessment Program: P-
13-42 Diatom Cleaning by Nitric Acid Digestion with a Microwave Apparatus. The
Academy of Natural Sciences. Report No. 02-06.
American Public Health Association, American Waterworks Association, Water Environment
Federation (APHA, AWWA, WEF). 1992. Standard Methods for the Examination of
Water and Wastewater. 18th ed.
Jan Stevenson, personal communication Aug. 11, 2009.
3.4 Preparation of Diatom Slides using Naphrax™ Mounting
Medium
Accurate identification and enumeration of diatoms requires mounting of cleaned material
between a microscope slide and coverslip in a medium with a high refractive index so that the
features of diatom frustules or valves are clearly visible at high magnification. Naphrax™ is a
commercially-available toluene-based mounting medium with high refractive index. This protocol
details the steps necessary to produce high-quality diatom mounts from cleaned diatom
material.
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3.4.1 Safety Precautions
Naphrax™ should be considered a hazardous substance because it contains toluene, an
organic solvent. Toluene volatilizes readily when heated. For this reason, the heating of
Naphrax™ should only be performed under a positive-draw fume hood. Personnel should wear
safety glasses and protective hand wear when working with liquid Naphrax™ at room
temperature, when heating Naphrax™ in a hood, or when in contact with solidified Naphrax™
toward the final stages of slide preparation.
Hot plate temperatures required for this procedure are high enough to cause severe burning of
exposed skin. Use extreme care when manipulating slides on the hot plate and when working
close to the hot plate.
3.4.2 Recommended Apparatus/ Equipment
• Corning ceramic-top hot plate with temperature control
• Positive-draw chemical hood
• Aluminum drying plate (25.5 x 20.0 x 0.5 cm, solid aluminum; lines forming 48 squares,
each 3.2 cm on a side, are etched on the surface. Each square is etched with an
identifying number)
• Glass microscope slides (1x3 inches; 2.5 x 7.5 cm)
• Glass coverslips (18 mm x 18 mm or 22 mm x 22 mm) - No. 1 thickness, stored in covered
glass jar filled with 100 % ethanol.
• Naphrax™ mounting medium
• Diamond scribe
• Disposable plastic pipettes
Adjustable pipettor (0 - 250ul); adjustable pipettor (200 - 1000ul)
• Pipette tips for adjustable pipettors
• Round-style tooth picks
• Forceps
• Polished, rounded wooden splints
• Wash bottle filled with distilled (DW) or reverse osmosis (RO) water
• Single-edged razor blades
Ethanol, 70%
• Acetone
• Kimwipe® tissues
• Slide labels
• Wax (the kind commonly used for candle making and canning foods)
10% HCI
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3.4.3 Methods
Estimate the amount of cleaned diatom material to deposit on coverslip.
1. Starting with cleaned material contained within 20-ml glass vials, estimate the volume of
suspended material that will need to be deposited ("dripped") on a coverslip to produce a
slide of the appropriate cell density. The ideal density to be achieved on the final mount is
somewhat subjective and is based on the amount of debris in the sample, the preferences of
the slide analyst, and the way in which the slide is to be used (e.g., counting,
documentation). Generally, between 10 and 20 diatom specimens should be present in a
single high power microscope field (1000X). The number of specimens per field will need to
be reduced if samples contain considerable amounts of silt. This estimate is referred to as
the "drip count" (the amount of cleaned material to be placed on a coverslip). Accuracy of
estimates improves with experience. In many cases, analysts will request that both a
"heavy" slide (-40 cells/field at 400 - 450x magnification) and a "light" slide (-30 cells/field
at 400 - 450x magnification) be made. When slidemaking is complete, record the final
amounts dripped on the Diatom Enumeration Data Sheet. Also note observations of
interfering materials (sand, silt, etc.) on this form.
2. In some cases, the number of diatoms in a sample is very sparse. This is usually because
diatoms were rare in the habitats sampled or the sample bottles contain a small amount of
material. In these cases, additional procedures (steps 3 to 5 below) are required to either
make a satisfactory slide for analysis or to determine that analysis of a sample is not
practical.
a. If a satisfactory slide could be made by increasing the concentration of cleaned
diatom material by two to five times, then do this by using a micropipettor to remove
the required amount of water from the vial of material after it has been allowed to
settle for at least eight hours. Spin the vial carefully while still resting on the table top
early, after about 4 h of the 8 hour period, to dissociate diatoms that have adsorbed
onto the side of the vial so they will settle. Record the concentration factor on the
Diatom Enumeration Data Sheet.
b. If a concentration of cleaned material greater than two to five times is required, then
re-subsample the original sample (Section 3.2). Take a subsample of a size sufficient
to prepare satisfactory slides. Use the entire remaining sample only if absolutely
necessary. Digest the subsample and prepare a new vial of cleaned material
(Section 3.3). Repeat step 1, above. If the concentration of cleaned material is still
not sufficient, concentrate it, as described above. If still too dilute, combine the two
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vials of cleaned subsample materials. Record steps, volumes, and final
concentration factor on the Diatom Enumeration Data Sheet.
c. If, after following the steps above to concentrate the cleaned material, the density of
diatoms on a coverslip still does not meet the criteria of 30 to 40 cells per field at 400
- 450x magnification, proceed to make the densest slide possible and take it to a
diatom analyst to evaluate. The analyst will make a determination of whether it is
practical to analyze the sample. He/she will quickly scan the slide in its entirety under
100x magnification, and estimate the total number of individuals on the slide. Then
he/she will make his/her determination of whether the slide is countable, taking into
account the density of diatoms, evidence of dissolution, and amount of debris (silt,
clay, broken remains of diatoms and other siliceous organisms) that would make it
difficult to identify specimens accurately. As a general guideline, if accurate
identifications are possible, and at least 100 specimens could be counted within four
hours, he/she should determine that the slide be analyzed; otherwise it should not. If
the diatom analyst determines that the slide should not be counted, inform the
Logistic Coordinator at 231-941-2230 immediately. Only under very special
circumstances will an analyst be asked to take the extraordinary measure of counting
a slide for a very long time (more than four hours). Record results of the diatom
analyst's determination and rationale on the Diatom Enumeration Data Sheet.
d. When doing an evaluation of a slide with few diatoms, as described in the step
above, a diatom analyst may occasionally see evidence suggesting that a sample
contains lightly silicified diatoms that may not have survived the digestion process. In
these rare instances, he/she may suggest that a "burn mount" be made to determine
whether diatoms did exist in the original sample. (This is one reason why a small
portion of the initial sample should always be saved, even for phytoplankton.) The
burn mount procedure was used extensively to create slides for diatom analysis
before the introduction of methods incorporating acids for the digestion of organic
material. Even though this method does not rid sample material entirely of organic
debris, diatoms on the slide can at least be identified as diatoms. For this method,
follow the EPA (1973) procedure. Briefly, a known portion of the untreated sample is
dripped onto a coverslip and allowed to dry at room temperature. When the sample
is dry, it is placed onto a hot plate and left for about 30 min at approximately 570°C.
The coverslip is mounted according to step 7, below. After it is prepared, have the
burn mount slide examined by a diatom analyst. He/she will determine if diatoms are
present and whether analysis of the slide is warranted. Slides prepared using the
burn mount method cannot be counted if too much organic material remains on the
slide. This is because it is not possible to make accurate taxonomic identifications.
Generally, burn mounts are used only as a last resort, and to confirm that weakly
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silicified diatoms are not present in the sample. Record information on all burn mount
attempts, successful or unsuccessful, on the Diatom Enumeration Data Sheet.
Include date, name of preparer, volume of subsample used, and whether diatoms
were observed.
Deposit cleaned material on coverslip.
3. Use forceps to remove single coverslips from the storage container and carefully clean each
by wiping with a Kimwipe®. Place each coverslip on a marked space. Be sure the drying
area is clean and dry to avoid cross-contamination. If the intended drip count will be less
than 600ul, drip an amount of distilled water onto the coverslip with a disposable pipette that
will be sufficient to form a thin layer of water over the entire coverslip when the diatom
suspension is added. Agitate the sample vial to homogenize the diatom suspension. Using
an adjustable pipettor, quickly withdraw the required amount from near the central portion of
the sample. Eject this material smoothly and carefully onto the layer of distilled water already
on the slip. By alternately drawing material up into the pipette and ejecting it, a
homogeneous suspension is achieved on the coverslip. In the case where more than ~600ul
of original sample is required, the addition of distilled water is not necessary, and the sample
can be ejected and mixed directly on the coverslip. In both cases, take care to ensure that
the suspension covers the entire surface of the coverslip, including the extreme edges of the
corners. Add 10% HCI to the cleared slurry (2 drops per 20ml of material in vial) to achieve
a more even distribution on the cover slip.
4. Should the coverslip overflow, discard the coverslip, wipe the drying area, and repeat the
procedure with a freshly cleaned coverslip. Discard the pipette tip when finished with each
sample. Once the marked space is loaded with coverslip preparations, it should remain
undisturbed until the coverslips are dry. At this point, drying of the slips can proceed at room
temperature (a period of several hours will be required), or gentle heat (warm to the touch
only) may be applied to hasten evaporation (a crook-neck lamp with incandescent light bulb
placed 15 - 30 cm over the coverslips is one option or a slide maker provides heat).
However, rapid evaporation with heating can produce strong patterns of diatoms settling on
coverglasses, which should be avoided. Once completely dry, put the coverslips on the hot
plate that has been preheated to 120 to 150°F. Leave for three to five minutes. This
procedure ensures that nearly all water is driven from the material on the coverslips and
helps assure that the diatom frustules will adhere to the surface of the glass. Remove the
coverslips from the hotplate and inspect them. If the pattern of diatoms distributed on any of
the coverslips is not even and smooth, they should be re-dripped. If coverslip distributions
seem unsatisfactory after repeated attempts, consult an algal analyst.
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Mount coverslip on microscope slide.
5. Using a diamond scribe, etch microscope slides with Sample ID, Subsample ID and Slide
Replicate ID.
6. Mount coverslip on slide using the following steps.
THE FOLLOWING STEPS MUST BE PERFORMED IN A POSITIVE-DRAW FUME HOOD!
7. Using a rounded wooden splint or disposable pipette, transfer a small amount of Naphrax™
(volume equivalent to two to four drops of water) to the central portion of the etched side of
the microscope slide. Then remove the appropriate coverslip from the aluminum plate with
forceps, being careful to handle the coverslip only at the extreme corners. Invert the slip and
place it gently on the Naphrax™ covered portion of the slide. Then place the slide (coverslip
up) on the hotplate and apply gentle heat until the evolution of bubbles resulting from the
evaporation of the toluene solvent first occurs, and then significantly diminishes. Heating the
slide will allow the Naphrax to flow under the coverslip. Remove the slide from the hot plate,
and, using the rounded toothpicks, gently position the coverslip and press it to form a
uniform, thin layer of Naphrax™ beneath the entire coverslip. Make sure that the edges of
the coverslip are brought parallel to the edges of the microscope slide. Care must be taken
at this stage not to press so hard as to damage or dislodge the diatoms or cause warping of
the coverslip. As this procedure is taking place, the Naphrax™ is "setting up" (becoming
hard), and the ability to move the coverslip will diminish rapidly. At this point, set aside the
mount to finish cooling.
8. Use a single-edge razor blade to carefully trim any excess Naphrax™ which has been
squeezed out from beneath the coverslip. Great care must be taken to avoid "lifting" the
coverslip by inadvertently allowing the edge of the blade to move between the coverslip and
the microscope slide. Once most of the excess Naphrax™ has been removed and
discarded, and while still working under the hood, place the mount in successive baths of
acetone, and then ethanol for no more than 10 or 15 seconds each. Finally, wipe the mount
clean with a Kimwipe® tissue.
9. Add a paper label to slides before slides have been analyzed.
10. Assemble forms and transmit slides. Put slides in plastic slide boxes; label each with name
of project and subproject, Subproject ID, box_ of _, date (month/year) box prepared, and
name or initials of preparer.
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11. Preserve and store cleaned material. After slides are analyzed according to the appropriate
protocol, and no additional slides need to be made, process the vials containing the
remaining acid-cleaned material for long-term storage. Working under a fume hood, add two
to four drops of 100% buffered formalin to each vial (some contractors use alcohol as a
preservative instead). Adding glass beads to the vials will also help to preserve the diatoms.
Tightly cap the vials and seal them by immersing the top 1/3 of the vial in melted wax. Then
transfer the vials to the appropriate storage cabinet for long-term storage. Be sure that the
cabinet and shelves on which they are stored are properly labeled with the study name,
year, and Subproject ID.
3.4.4 Quality Assurance/ Quality Control
It should be understood that, given the microscopic size and large numbers of diatoms which
are transferred from the cleaned material vials to the finished mount, there are a number of
steps where contamination of the samples is possible. Laboratory rooms where raw or
processed samples are handled should be kept as clean as possible. Laboratory bench
surfaces should be kept clean and free of debris at all times. Techniques similar to those used
for sterile experiments (bacteriological plating, etc.) should be followed to minimize the risk of
cross-contamination of samples. All equipment coming into contact with sample material should
be rinsed in DW or RO water at least three times. Disposable pipettes should be used when
possible.
The distribution of specimens on the final mounted coverslips should represent the samples
contained within the cleaned material vials. The degree to which this is true depends on how
well the cleaned material is dispersed prior to subsample withdrawal, and how evenly the
withdrawn material is dispersed on the coverslip. Great care should be taken to ensure that
these two steps are completed properly.
3.4.5 References
American Public Health Association, American Waterworks Association, Water Environment
Federation (APHA, AWWA, WEF). 1992. Standard Methods for the Examination of
Water and Wastewater. 18th ed.
Patrick, R. and C. Reimer. 1967. The Diatoms of the United States. Vol. 1. Monograph No. 13,
Academy of Natural Sciences of Philadelphia, 688 pp.
PCER, ANSP. 1990. Laboratory Safety Manual.
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USEPA. 1973. Biological Field and Laboratory Methods for Measuring the Quality of Surface
Waters and Effluents. EPA-670/4-73-001. US EPA Office of Research and
Development. Cincinnati, OH.
3.5 Analysis of Diatoms
The purpose of the diatom subsample analysis is to estimate the proportion of diatom taxa
found in a count of 600 valves (one-half of an individual diatom cell). On average, analysis of a
slide should take approximately two hours; in no case should it exceed four hours or six
transects. Biovolume measurements and imaging are part of this count process and should be
completed within the four hour time frame. This does not include time spent learning new taxa
when analyzing the first few samples in a new study unit. Images should be taken of all taxa
encountered, including unknowns. Images need to be of sufficient quality as to be used for
identification. Photos should be taken of all taxa listed on the taxa list over all of the samples.
3.5.1 Recommended Apparatus/ Equipment
• Compound microscope
• Oil immersion objective (100x) with a numerical aperture of at least 1.3
• Eyepieces of 10-15x
• DIG (differential interference contrast) or bright field condenser
• Diamond scribe mounted on microscope's objective stage
• High intensity light source
3.5.2 Methods
3.5.2.1 Diatom counts.
1. Scan slides at low to medium magnification (100x to 450x) to confirm that diatoms are
evenly distributed on the coverslip and are at a density appropriate for efficient counting. At
high magnification (1000x), there should be 10-20 diatoms per field. If there are problems
with dispersion or density that would compromise the quality and accuracy of the analysis,
discuss these with Diatom Preparation Lab personnel and have new slides made. Avoid
counting diatoms in any disrupted areas of the mount, particularly edges that have optical
aberrations. If diatoms on the slides are very sparse, refer to procedures in Section 3.4 for
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handling low-density samples. Always save any count data generated for a sample, even if
the number of valves orfrustules is low (e.g., <100).
2. Because slides may need to be recounted for QA/QC purposes, it is very important to clearly
demarcate the areas of a slide scanned during a count. After the preliminary slide
examination, secure the slide in the mechanical stage and use the microscope's diamond
scribe to etch a horizontal or vertical line (depending on personal preference) on the
coverslip to mark the edge of the first row to be counted. Etch at least three lines on the
diatom slide (if etching before counting) or a line for each row counted (if etched during an
analysis). Rows are narrow rectangular areas (strips) of the slide adjacent to the scribed
line, with width equal to the field of view. Start rows far enough from the coverslip edge to
avoid optical distortion, and end them near the opposite coverslip edge where diatoms are
no longer clearly visible (see diagram below). Locate a starting point near one end of the
etched line and make a circle with the scribe. This denotes the starting point of the count.
During the count, etch a circle around the last field counted in the first row and at the
beginning and end of all other rows. Always check to make sure that etching is clearly
visible so that circles and lines can be located easily by others.
3. When the line and first field are etched on the coverslip, and the first field is focused under
oil immersion, begin the count. Record the transect start coordinates from the microscope
stage.
4. Count diatom valves along transects, which are narrow rectangular areas (strips) of the slide
with width equal to the field of view. The transect width should be the maximum width
common to all participating taxonomists (at least 90 microns)
5. Count 600 valves. Count all partial valves that are more than 50% of the valve or that
contain unique features such as recognizable central areas or distinct ends. Count all valves
and fragments that extend at least halfway into the transect. Record the transect end
coordinates from the microscope stage.
3.5.2.2 Biovolume measurements.
Biovolumes should be determined for all taxa either by measurements of cells observed in the
project samples, from databases with known sources, or from the literature. The following
criteria should apply regardless of the source of measurements.
Biovolumes for each abundant taxon (i.e., occurring in more that 5% in any one sample) should
be based on measurements of 10 cells or more.
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Biovolumes for each common taxon (i.e., occurring 2-5% in any one sample) should be based
on measurements of one cell or more.
Biovolumes for each rare taxon (i.e., occurring in 0.1-2% in any one sample) should be based
on measurements from literature descriptions of taxa or measurements of one cell or more.
For each taxon requiring biovolume measurements, select a simple geometric figure matching
the shape of the taxon as best as possible, and determine the dimensions that must be
measured. Average biovolume for a taxon should be calculated as the average of biovolumes
of cells measured versus the average dimensions of cells measured. When a dimension of a
cell is not known (e.g. depth), it should be estimated based on sizes of cells of the same taxon
with similar known dimensions. Depth can be estimated by focusing on tops and bottoms of
cells if other cells of the same taxon can not be found for measure of unknown dimensions.
The calculation of phytoplankton abundance depends on the apparatus used during analysis.
Biovolume values are determined by multiplying the abundance (cells/ml) by the average
biovolume of each cell (urn3). Phytoplankton abundance (cells/ml) is calculated as follows:
„ . , count* chamber area* sample volume* subsampleDCF
cells/ ml — ——
trans _lengthx trans _width* chamber_volume
where count = number of cells counted, fieldDCF = field dilution/concentration factor, chamber =
chamber area (in mm2), subsampleDCF = subsample dilution/concentration factor, field =
microscope field area (in mm2), numfields = number of microscope fields, and chambervol =
chamber volume used.
3.5.3 Quality Assurance/ Quality Control
Sample and slide quality can affect the outcome of these procedures. Minor deviations that do
not affect the area scanned or number of specimens observed should be described on bench
sheets. Other deviations should be discussed with the Project Manager for inclusion in the
project QA/QC notes. A total of 10% of the samples collected from each study unit will be
analyzed for quality control. EPA will identify which random slides will be sent to an
independent laboratory for re-counting. Refer to Section 3.10.
3.5.4 References
Cotter, P. 2002. "Tabulator" Installation and User's Guide. Version 3.51. ANSP, PCER.
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Hillebrand, H., C. D. Durselen, D. Kirschtel, U. Pollingher, and T. Zohary. 1999. Biovolume of
pelagic and benthic microalgae. Journal of Phycology 35:403-242.
Moulton, S.R., II, J.G. Kennen, R.M. Goldstein, J.A. Hambrook. 2002. Revised protocols
for sampling algal, invertebrate, and fish communities in the National Water-Quality
Assessment program, U.S. Geological Survey Open-File Report 02-150. In Press.
PCER, ANSP. 2002. Diatom Cleaning by Nitric Acid Digestion with a Microwave
Apparatus. Protocol No. P-13-42.
PCER, ANSP. 2002. Analysis of Soft Algae and Enumeration of Total Number of Diatoms
in USGS NAWQA Program Quantitative Targeted-habitat (RTH and DTH) Samples.
Protocol P-13-63.
PCER, ANSP. 2002. Preparation of Diatom Slides Using Naphrax™ Mounting Medium.
Protocol No. P-13-49.
Porter, S.D. 1994. Amendment to Guidance, Procedures, and Specifications for
Processing NAWQA Algal Samples by Contract Laboratories. Email contract reference
of 10/19/1994 from Stephen Porter to Allison Brigham of the USGS, Ann St. Armand of
Phycotech, Don Charles and Frank Acker of the Academy of Natural Sciences.
Porter, S.D., T.F. Cuffney, M.E. Gurtz, M.R. Meador. 1993. Methods for Collecting Algal
Samples as Part of the National Water Quality Assessment Program. U.S. Geological
Survey Open-File Report 93-409, Raleigh, NC. 39 pp.
http://water.usgs.gov/nawqa home.html..
United States Geological Survey, National Water-Quality Assessment Program. 1997.
Protocols for Processing NAWQA Algal Samples. Draft Manuscript. February 1997.
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3.6 Analysis of All Algae
This protocol describes a quantitative procedure for analyzing all algae in samples. This
procedure is semi-quantitative (i.e. describing cells per sample, per unit substrate area sampled,
or per unit water volume sampled) and designed to provide data on algal densities and amount
of algal biovolume. This protocol describes procedures for identification, counting,
documentation, and measurement of algae. Often this protocol is referred to as the soft-algal
count, which refers to the emphasis on identification of non-diatom algae.
This protocol also describes a tiered analysis of all algae. The simplest protocol is the 300
natural unit count, which was used in NAWQA, EMAP, and NLA programs.
With the exception of time spent learning new floras, the counting of 300 natural units should be
finished in approximately two hours, on average.
3.6.1 Definitions
Natural counting unit. Algae grow unicellularly or in multicellular filaments and colonies. Each
natural grouping of algae (i.e., each individual filament, colony, or isolated cell) is defined as a
natural counting unit. Diatoms are an exception; each diatom cell is always considered a natural
counting unit, even if attached to other cells. The main purpose of using 'natural counting units'
is to prevent a colonial or filamentous form from dominating a count. It also facilitates the
counting of algal forms which have linked cells that may be hard to distinguish.
Palmer-Maloney Count. This count type is to be completed on all samples. A maximum
volume of 0.1 ml of the soft algal subsample viewed in two Palmer-Maloney cells will be
assessed.
Using a transect approach, soft algae and live diatoms are enumerated until 300 natural
counting units are reached or a total transect length of 115.7 mm (for field of view 0.54 mm) is
assessed. Dead diatoms are also enumerated.
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Dilution liquid. Samples collected as part of many sampling programs, including NAWQA, are
preserved with a 2-3% formalin solution. Dilution liquid refers to the 3-5% formalin solution
added to fractions when it is necessary to make dilutions of these fractions.
Macroalgae. A soft-algae component that consists of visible colonies or filaments.
Microalgae. A soft-algae component that consists of forms which are not visible without the aid
of a microscope.
3.6.2 Recommended Apparatus/ Equipment
Compound microscope with 20x objective for a total system magnification of 200x and 40-
45x objectives for a total system magnification of 400-450x.
Glass microscope coverslips, rectangular, 26 x 60 mm, #1 thickness.
Glass, wide-bore pipettes > 1 mm inside diameter, or eye-dropper.
Two Palmer-Maloney Counting Cells with ceramic chamber (chamber depth of 0.4 mm;
volume of 0.1 ml) or metal chambers
Glass pasteur pipettes, 5.25 inch, < 1 mm diameter.
Glycerin
3.6.3 Methods
Palmer-Maloney count
1. Place a rectangular cover slip (#1 thickness, 22 x 50 mm) at 45° to the counting cell,
covering about 1/3rd of the chamber, but not across the center of the cell.
2. Thoroughly mix the Palmer-Maloney fraction and draw into the micro-pipette 100 ul (plus
suitable additional amount to allow for displacement from the chamber by the cover slip and
prevent the occurrence of air bubbles). Quickly add the fraction drop-wise, into the center of
the chamber. When the surface tension starts to draw the cover slip across the chamber,
adjust the sides of the cover slip so that ends of the chamber are covered and the cover slip
hangs over both sides of the ceramic portion of the counting cell.
3. Add glycerin to the area where the cover slip extends past the ceramic portion. This seals
the cover slip to the counting cell temporarily (without excess heat or vibration, the counting
cell can be used for a week or more).
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Palmer-Maloney Fractions.
4. Dilutions or concentrations of samples with extremely high or extremely low diatom
abundance may be necessary. This must be considered on a case by case basis. In
general 15-30 natural counting units (both diatom and soft algae) should be visible per
field of view. Assessment of five fields of view is advised. For samples where the number
exceeds 30 natural counting units, dilution is recommended. Where the number is < 15,
consider concentration. Silt/detritus levels are also a factor. In samples with high silt/detritus
levels, even if there are < 15 natural units per field of view, consider dilution. Avoid
concentrating samples were the silt/detritus level would be raised to a level too high to
complete a successful count. Note on the count sheet when silt/detritus levels prohibit
concentration or lead to a dilution. Both Palmer-Maloney cells should be prepared using the
same Palmer-Maloney fraction. The S-R count should be completed on the original sub-
sample taken from the total collected material. Only concentrate/dilute a portion (10 ml or
less) of the original sub-sample. No sample should be diluted or concentrated to more than
20 % of its original volume.
Note: For samples with high amounts of sand and/or silt, place the aliquot in the Palmer-
Maloney Cell for assessment. Dilute if necessary and recheck sample. If large inorganic
particles are present (but NOT organic detritus) and blocking view of algal cells, again
thoroughly mix the Palmer-Maloney fraction. Allow the sample to stand while slowly counting
to 5 or 6 (approximately 4-5 seconds). Re-load the Palmer-Maloney cell with the subsample
according to instructions and count the sample or dilute again as necessary. This procedure
is valid ONLY for samples with large amounts of inorganic sand and silt. No settling should
occur with samples that contain organic detritus or at any time during the dilution process.
Once the Palmer-Maloney cell is prepared, allow chamber to stand for up to 10 minutes to
allow contents to settle.
5. Palmer-Maloney fraction data should be recorded on a separate log and the sheet included
with the deliverables.
Enumerate 300 natural counting units.
6. Position the microscope stage so that the first field of view is in the top left hand area of the
Palmer-Maloney cell at, but not overlapping, the cell edge. Using transects, of varying
length, with a width of 0.54 mm (or width of the field of view) scan up to, but not exceeding,
a total transect length of 115.7 mm (or equivalent). Every other transect, moving down the
Palmer-Maloney cell, is to be enumerated, i.e. each transect enumerated should be
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separated by a width of one field of view. This will prevent field of view overlap. For
microscopes where the 40x field of view differs from 0.54 mm calculate the maximum
transect length required. The full length of each transect should be scanned for algal
enumeration. Record the start and end coordinates for each transect for the purpose of total
transect length calculation.
7. Identify and enumerate all soft algal forms within the field of view. Algal forms are
enumerated using algal natural units and identified to the lowest possible taxonomic level.
8. The taxa names to be used for this project, when identifying soft algal forms, are listed in the
NRSA 2008 start list for soft algae [amended at the 2008 NAWQA soft algal workshop to the
current accepted taxonomy]. This is to aid with taxonomic harmonization. This list helps
promote consistency in identifications among laboratories.
9. Diatoms are differentiated as to "living" or "dead" at the time of collection. If there is any
protoplast material in the frustules the diatom is considered to have been living when
collected.
10. Count the number of algal cells for each multicellular natural unit. When cell length within a
filament cannot be determined then a standard length of 10 urn per cell should be used.
Notes should be made on the count report when this standard length is employed.
11. The number of cells for multicellular natural units is recorded in parenthesis beside the tally
of natural counting units. All diatoms are grouped into one category - undifferentiated
diatoms.
12. Tally data onto a bench sheet or into a computer based program. Notes on difficulties
encountered in taxa identification due to the use of Lugol's should be made during the count
and logged with the count data.
13. Repeat procedures until 300 natural counting units have been enumerated. Only "living"
diatoms are counted against the needed 300 natural algal units. "Dead" diatoms are
recorded separately to inform the proportion of living vs dead diatoms in the sample.
14. Digital images are required for all identified taxa using the criteria outline in the Images
section of this document (Section 3.9.1).
15. Biovolume measurements are required for all identified taxa using the criteria outline in the
Biovolumes section of this document (Section 3.6.4).
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16. Once 300 natural counting units have been enumerated, or the total transect length has
been assessed, cease counting and record the total length of transects scanned.
3.6.4 Calculations and Reporting
The calculation of algal cell density and biovolume can be made per sample, per unit area of
substratum sampled, or per volume of water sampled for qualitative and quantitative analyses of
benthic and planktonic algae. Cell abundance per sample (cells/sample) is calculated as
follows:
Equation 1
count x chamber _area x sample _volume x subsampleD CF
cells I sample =
trans _length x trans _width x chamber _volume
where count = number of cells counted, chamber_area = chamber area (in mm2),
sample_volume=tota\ volume (ml) of sample from which subsamples were drawn,
subsampleDCF = subsample dilution/concentration factor, transjength and trans_width are the
length (mm) and width (mm) of transects counted, and chamber_volume = chamber volume
used.
The subsampleDCF = subsample dilution/concentration factor is calculated by dividing the final
diluted or concentrated subsample volume by the initial volume. So, a 10 ml_ subsample
concentrated to 1 ml_ would have a subsampleDCF =0.1. A 10 ml_ subsample diluted to 100
ml_ would have a subsampleDCF=10.0.
Currently, there are not plans to calculate cells per unit area or volume of samples collected in
the NWCA. If there were, cells per unit area or volume sampled can be determined by dividing
the cells/sample by the area or volume sampled. If the laboratory sample (i.e. sample_volume)
is a subsample of a larger sample collected in the field, then the laboratory sample fraction of
the total field sample should be taken into account to determine the area or volume sampled
corresponding to the laboratory sample.
Biovolume values are determined by multiplying the abundance (e.g. cells/sample) by the
average biovolume of each taxon (urn3).
Reports should be presented in excel spread sheets or comma delimited files. A template will be
provided that includes rows for: sample ID, site ID, site visit number, date sampled, full taxon
name, number of cells of the taxon counted, number of cells of the taxon per sample, and
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biovolume of the taxon counted. Divide number of valves by two to calculate diatom cell
abundance. Separate files should be prepared for the diatom and all-algae counts.
3.6.5 Quality Assurance/ Quality Control
It is critical that prior to taking a small portion of the subsample, the sample be thoroughly
mixed. A small portion of the subsamples from each group of samples is re-processed using
final dilution/concentration factors and checked for number of natural units per microscope field.
Minor deviations to this protocol are expected, especially concerning methods to break up algal
clumps. These deviations should be reported to project managers and/or algal taxonomists and
noted on the Data Form. About 10% of the sample vials will be swapped and sent to an
independent laboratory for a QA/QC analysis. Refer to Section 3.10.
3.6.6 References
Palmer, C.M. and I.E. Maloney. 1954. A new counting slide for nannoplankton. American
Society of Limnology and Oceanography Special Publication Number 21. 6 pp.
PCER, ANSP. 2002. Analysis of Diatoms in USGS NAWQA Program Quantitative Targeted-
Habitat (RTH and DTH) Samples. Protocol No. P-13-39.
USEPA. 2008. National Rivers and Streams Assessment: Laboratory Methods Manual. EPA
841-B-07-010. U.S. Environmental Protection Agency, Office of Water and Office of
Research and Development, Washington, DC. Protocols for NRSA 2008-2010 revised
2-26-09.
United States Geological Survey, National Water-Quality Assessment Program. 1997.
Procedures for Processing NAWQA Algal Samples. Draft Manuscript. February 1997.
Weber, C.I. 1973. Biological Field and Laboratory Methods for Measuring the Quality of
Surface Waters and Effluents. EPA-670/4-73-001. National Environmental Research
Center, Office of Research & Development, U. S. Environmental Protection Agency.
Cincinnati, OH.
3.7 Phytoplankton Chlorophyll a
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Chlorophyll-a samples are filtered in the field, placed in a labeled vial, and stored on ice until
arrival at the laboratory. The laboratory should store the filter in the freezer at -20°C ±2°C for
no more than 3 1/4 weeks before analysis.
In addition to specifying laboratory methods for sample analysis, we use a performance-based
approach that defines a set of laboratory method performance requirements for data quality.
Method performance requirements for this project identify detection limit, precision, and
accuracy objectives for each parameter (Table 3.7-1).
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Table 3.7-1. Performance Requirements for Chlorophyll a Analytical Methods.
Analyte
chlorophyll a
Units
ug/L (in
extract)
Potential Range
of Samples
0.7 to 11, 000
Long -Term
MDL
Objective2
0.5
Laboratory
Reporting Transition
Limit3 Value4
1.0 15
Precision
Objective5
±0.5 or ±10%
Bias
Objective6
±0.5 or ±10%
1 Estimated from samples analyzed at the WED-Corvallis laboratory between 1999 and 2005 for TIME, EMAP-West, and WSA streams from across the U.S.
2 The long-term method detection limit is determined (eq. 1a) as a one-sided 99% confidence interval from repeated measurements of a low-level standard
across several calibration curves, and includes medium or mean method blank results, (USGS Open File Report 99-193, EPA 2004). These represent
values that should be achievable by multiple labs analyzing samples over extended periods with comparable (but not necessarily identical) methods.
3 The minimum reporting limit is the lowest value that needs to be quantified (as opposed to just detected), and represents the value of the lowest nonzero
calibration standard used. It is set to 2x the long-term detection limit/fractional spike recovery, following USGS Open File Report 99-193 and EPA 2004.
4 Value at which performance objectives for precision and bias switch from absolute (< transition value) to relative 9> transition value). Two-tiered approach
based on Hunt, D.T.E. and A.L. Wilson. 1986. The Chemical Analysis of Water: General Principles and Techniques. 2nd ed.. Royal Society of Chemistry,
London, England.
5 For duplicate samples, precision is estimated as the pooled standard deviation (calculated as the root-mean square) of all samples at the lower
concentration range, and as the pooled percent relative standard deviation of all samples at the higher concentration range. For standard samples,
precision is estimated as the standard deviation of repeated measurements across batches at the lower concentration range, and as percent relative
standard deviation of repeated measurements across batches at the higher concentration range.
6 Bias (systematic error) is estimated as the difference between the mean measured value and the target value of a performance evaluation and/or internal
reference samples at the lower concentration range measured across sample batches, and as the percent difference at the higher concentration range.
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3.7.1 Summary of Method
Chl-a containing phytoplankton in a measured volume of sample water, are concentrated by
filtering at low vacuum through a glass microfiber filter under low light conditions. The filter is
then placed into a labeled centrifuge tube and frozen at -20° C. The pigments are extracted from
the phytoplankton in 90% acetone with the aid of a mechanical tissue grinder. The filter slurry is
allowed to steep for a minimum of two hours to ensure thorough extraction of the chl-a. The
sample is centrifuged for 15 minutes at 675 x gravity. An aliquot of the supernatant is
transferred to a glass cuvette and fluorescence is measured on the Turner Fluorometer. The
concentration of chl-a in samples is determined from the calibrated instrument reading and from
dilutions as necessary. The concentration of chl-a in the natural water sample is reported in
ug/L. This SOP also includes chl-a standard preparation and calibration of the fluorometer.
3.7.2 Definitions
Laboratory Blank: An aliquot of reagent water that is treated exactly as a sample including
exposure to all glassware, equipment, solvents, reagents, and surrogates that are used with
samples. The laboratory blank is used to determine if analytes or interferences are present in
the laboratory environment, the reagents, or the apparatus.
Primary Calibration Standard: A solution containing an analyte that is prepared using a
reference material traceable to EPA, the National Institute of Standards and Technology (NIST),
or a source that will attest to the purity and authenticity of the reference material.
RO water: Water passed through a reverse osmosis system.
SSCS: Secondary Source Check Standard is a sample containing the analyte of interest at
known concentrations. The SSCS is obtained from a source external to the laboratory or is
prepared from standards obtained from a different source than the calibration standards. The
purpose is to check laboratory performance using test materials prepared independently from
the normal preparation process.
3.7.3 Interferences
After the filters are ground the samples are cloudy and filter particles are suspended in solution.
The samples must be clarified by centrifugation prior to analysis to prevent interference.
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Contamination that fluoresces in the red region of the light spectrum may interfere in the
accurate measurement of chl-a.
Interference by chlorophylls b and c are avoided by the use of the Turner Optical Filter Kit 10-
040. Thus, acidification of the sample followed by measuring fluorescence of the acidified
sample is not necessary for this method.
Minimum sensitivity settings on the fluorometer should be avoided due to quenching effects in
highly concentrated solutions. Dilutions should be performed instead.
Fluorescence is temperature dependent with higher sensitivity occurring at lower temperatures.
All standards and samples must be at the same relative temperature during analysis.
Photosynthetic pigments are light and temperature sensitive. All work must be conducted in low
light and samples must be stored in the dark at -20° C to prevent degradation.
3.7.4 Safety Precautions
This method does not address all safety issues associated with its use. The laboratory staff is
responsible for safely conducting lab work and chemical analysis in accordance with the
NHEERL-WED Health and Safety Handbook, the applicable Dynamac Hazardous Activity
Safety Plans, and the Material Safety Data Sheets for the specific chemicals.
Proper personal protection equipment (PPE) is worn to prevent exposure to organic solvents.
Measures need to be taken to minimize exposure to acetone fumes. Proper room ventilation
and use of a chemical fume hood is required for all procedures involving acetone. Consult the
MSDS for detailed safety and handling information for acetone
3.7.5 Equipment and Supplies
• Model 10-AU Digital Turner Fluorometer equipped with: 1) 13-mm cuvette holder; 2)
Optical Filter Kit PN 10-040, which includes: Blue lamp (PN 10-089); Emission Filter (PN
10-115); (436 nm) Excitation Filter (PN 10-113); (1 ND) Reference Filter (PN 10-032)
• Centrifuge, capable of 675 x gravity
• Tissue grinder, PowerGen 125
• Whatman GF/F glass fiber filters
• Centrifuge tubes, polypropylene, 50-mL capacity with screw caps
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• Tweezers or flat tipped forceps
• Vacuum pump capable of maintaining a vacuum of 6 in. Hg (20 kPa)
• Room thermometer
• 13 x 100 mm Borosilicate glass culture tubes (10 ml_)
• 10-mL pipette and disposable tips
• Graduated cylinders (various sizes as needed)
• Safety glasses
• Nitrile gloves
• Lab coat
• Laboratory exhaust fume hood
• Analytical balance with resolution to 0.01 mg
• Filter apparatus
• 50-mL volumetric flasks
• Explosion proof refrigerator
-20° C Freezer
• Spectrophotometer, UV-visible, located in EPA Main Building Room 246.
3.7.6 Reagents and Standards
Reagents
1. Acetone, reagent grade
2. Chlorophyll a free of chlorophyll b. (Sigma-Aldrich; P.O. Box 14508; St. Louis, MO 63178;
800-325-5832. C6144 from algae, C5753 from spinach. Purchased yearly.)
3. RO Water: Water passed through a reverse-osmosis system to produce water similar to
ASTM Type I reagent with 16.7 megaohms resistivity (Reference 16.5).
4. Primary and Secondary Calibration Standard Preparation
5. Gently tap the contents of the chl-a standard ampoule/vial to the bottom of the ampoule/vial.
6. Break or open the ampoule/vial. Transfer 1-3 mL of 90% acetone to the ampoule/vial. Using
a Pasteur pipette, carefully transfer the contents to a 50-mL volumetric flask. All glassware
must be clean and acid-free before use. Thoroughly rinse the ampoule/vial (including sides)
a second, then a third time with 90% acetone transferring the contents each time to the 50-
ml volumetric flask.
7. Fill the volumetric flask to volume using 90% acetone.
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8. Measure the chl-a primary standard using a UV-visible spectrophotometer. Verify the
measured value of chl-a against its calculated concentration.
9. After measuring absorbance at 750, 664, 647, and 630 nm, the concentration of chl-a in the
90% acetone standard may be calculated by the following equation, which corrects for
absorbance at 750 nm (simplified):
chl-a (mg/L) = 11.85 (Abs 664) - 1.54 (Abs 647) - 0.08 (Abs 630) - 13.31 (Abs 750)
10. Remake the standard if the chl-a concentration is outside 5% of expected value. If this is not
feasible, perform repeat measurements until a stock standard concentration can be
confidently assigned.
11. Prepare the secondary source check standard (SSCS) in the same manner as above using
an alternate source of chl-a (see Section 7.1.2).
12. Store all stock solutions in an explosion proof refrigerator at 4° C in a dark container.
Caution: Acetone is hazardous to your health and is a highly flammable material. Do not allow
skin contact or inhale vapors. Acetone rapidly degrades PVC. Wear gloves constructed of
resistant material and work in a chemical fume hood.
Preparation of Working Standards
1. Prepare working standards of chl-a using the primary standard solution. The dilutions are
prepared in 90% acetone to the final concentrations of 0, 2, 20, 50, 100 and 200 uL chl a/L.
2. Working standards are stored in an explosion proof refrigerator at 4° C in a dark container.
3.7.7 Sample Collection, Preservation and Storage
1. Filtering should be performed in subdued light as soon as possible after sampling. The
applied vacuum should be monitored with a gauge or manometer to ensure filtration
pressure does not exceed 6 in. Hg (20 kPa). Higher filtration pressures may damage cells
and result in a loss of chl-a.
2. Thoroughly shake the container to suspend particulates before the sample is poured for
filtering. Accurately measure 500 ml_ of sample. Remove the filter from the base with
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tweezers, fold once and insert inside screw cap centrifuge tube. Cover the outside of the
tube with foil for added protection from light.
3. Sample filters are stored frozen (-20° C) in the dark until extraction.
4. Samples can be stored up to 4 weeks before extracting (Reference 16.1).
3.7.8 Calibration and Standardization
Calibration of the Fluorometer
1. Allow the fluorometer to warm up for at least 15 minutes.
2. Calibrate the fluorometer using the 50 ug chl-a/L primary calibration standard. Make sure to
note the room temperature when recording calibration data.
3. Measure the fluorescence of each standard at the sensitivity setting that provides a
midscale reading.
4. Record all measurements and analyze the linearity of the curve to ensure r>0.999.
5. Calibration is performed before each analysis, or when there has been an adjustment made
to the instrument such as replacement of lamps, filters or the photomultiplier. The instrument
should also be recalibrated if the room temperature fluctuates ±3° C from the initial
calibration temperature.
Calibration Verification
1. Measure the 50 uL chl-a/L SSCS after calibration.
2. Record the SSCS measurement on the datasheet (Section 17.0).
3. Continue to measure the SSCS and a laboratory blank every 10 samples. If SSCS drifts
outside ±10% recovery, recalibrate.
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3.7.9 Procedure
Sample Preparation
1. Set up the filtration apparatus in the lowest light possible to prevent degradation of chl-a in
the samples. The filter apparatus should be clean and acid-free.
2. Set-up a standard laboratory vacuum apparatus using a trap to collect excess/overflow
water preventing aspiration into the vacuum system/pump.
3. Remove the samples from low-light containers and shake thoroughly to suspend the
particulates. Carefully measure a 500-mL aliquot of the sample, and filter it through a
Whatman GF/F glass microfiber filter. Vacuum filtration must not exceed 6 in. Hg (20 kPa).
Higher vacuums may damage cells and result in loss of chl-a. Important note: Don't allow
the filter to stay on the filter apparatus too long. The vacuum should be turned off at the
valve when the sample is filtered so as to protect the chl-a in the sample.
4. Remove the filter using forceps, fold it in half, and place into a centrifuge tube labeled with
an identification number. Centrifuge tubes should be placed in a lightproof secondary
container before storage.
5. Store the filter in a freezer at -20° C for at least 24 hours, to lyse the cells and release the
chl-a contained within them. The samples should be removed from the freezer and analyzed
within approximately 25 days.
Extraction of Filter Samples
1. Remove the samples from the freezer. Maintain low light throughout the extraction.
2. Pipette 40 ml_ of 90% acetone into each centrifuge tube.
3. The filters are ground completely using the PowerGen 125 tissue homogenizer. Rinse the
homogenizer thoroughly with RO water between samples to avoid cross-contamination.
Small pieces of filter remaining on the homogenizer need not be transferred into the sample
solution.
4. Vortex each tube to resuspend particulates within the solution.
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5. Samples should be allowed to steep between 2-24 hours in an explosion proof refrigerator
at 4° C.
6. After steeping is complete, vortex the sample again to resuspend particulates.
7. Centrifuge samples for 15 minutes at 675 x gravity. Allow samples to come to ambient
temperature before analysis. Check and record room temperature before analysis.
Caution: Acetone is hazardous to your health and is a highly flammable material. Do not allow
skin contact or inhale vapors. Acetone rapidly degrades PVC. Wear gloves constructed of
resistant material and work in a chemical fume hood.
Sample Analysis
1. Allow the fluorometer to warm up for at least 15 minutes.
2. Use a 90% acetone solution to zero the instrument on the sensitivity setting that will be used
for sample analysis.
3. Read the SSCS in the mid-range of the calibration that is pertinent to the sample range.
4. Transfer 8 ml_ of the sample into a borosilicate culture tube. Care should be used not to
disturb the solids at the bottom of the tube during the transfer.
5. Choose a sensitivity setting that yields a midscale reading when possible and avoid the
minimum sensitivity setting. If the concentration of chl-a in the sample is too high to be
measured, dilute the sample to the appropriate midscale range using 90% acetone.
6. Record the fluorescence measurement and dilution data if applicable on the data sheet (see
Section 17.0).
3.7.10 Data Analysis and Calculations
To calculate the concentration factor for chl-a, enter measurements into spreadsheet template,
which uses the equation:
Concentration factor = extraction solution volume (ml_)/sample volume (ml_)
Raw data are reported by electronic spreadsheet in ug/L, and this data is then used to calculate
chl-a using the following calculations:
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(1) Cf/a = (extraction solution volume/filtered volume)*(total volume/area scraped)
(2) Chl-a = raw data (ug/L) * Cf/a (ml_/cm2) * (1L/1000ml_)
Where:
Extraction solution volume = 40 ml_ (unless noted)
Filtered volume = volume of sample filtered (usually 25 ml_)
Total volume = volume of sample collected (usually 500 ml_)
Area scraped = (# of transects)*(area delimiter (cm2))
Cf/a = concentration factor per area (see above)
Duplicate precision is determined from analytical results.
Report results to three significant figures.
3.7.11 Method Performance
Method performance is measured through analysis of blank measurements and duplicate
precision. Blank measurements should be <0.015 ug/L. These measurements ensure accuracy
at low-level samples. Duplicate precision must be within 10%. On occasion duplicates will fall
outside this range; this variance is most likely a result of lack of homogeneity in the prefiltered
sample solution. The possible reasons for imprecise duplicate measurements include lack of
homogeneity, evaporation of acetone, and instrument drift, and should be noted on the
datasheet. If duplicates are not in good agreement and there is no apparent reason or
observable differences, the instrument should be recalibrated and samples reanalyzed.
3.7.12 Pollution Prevention
1. The chemicals used in this method pose little threat to the environment when properly
managed.
2. All standards and reagents should be prepared in volumes consistent with laboratory use to
minimize the volume of waste. Wastes are collected for recycling or appropriate disposal.
3.7.13 Waste Management
1. It is the laboratory's responsibility to comply with all federal, state and local regulations
governing waste management, and to protect the environment by minimizing and controlling
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all releases from fume hoods and bench operations. Compliance with all sewage discharge
permits and regulations is required.
2. Contact the Contractor Program Health and Safety Manager for guidance on waste
collection and disposal. For further information on waste management, consult The Waste
Management Manual for Laboratory Personnel, and Less is Better: Laboratory Chemical
Management for Waste Reduction, both available from the American Chemical Society's
Department of Government Relations and Science Policy, 1155 16th Street N.W.,
Washington, DC 20036.
3.7.14 Quality Control
Analytical Duplicate is a separate analysis from the same sample aliquot, run a minimum of
once every 10 samples. The analytical duplicate is the same aliquot run near the end of the run.
Blank is a reagent blank taken from the 90% acetone solution and measured as a sample. A
blank should be analyzed every 10 samples.
SSCS is mid-range in calibration and is measured every 10 samples. Recalibrate if outside 10%
of the expected value.
Room temperature should be monitored and the instrument recalibrated if the room temperature
varies ±3° C of the initial calibration.
Personnel performing this procedure are trained and must demonstrate their ability to handle
standards in normal laboratory conditions without significant photodegradation. Stock solutions
and working standards must routinely be transported through lighted hallways; during these
times they should be well protected against the fluorescent lighting, which is particularly harmful
to chlorophyll pigments. New analysts will demonstrate the ability to adequately light-proof
standards before transporting them through lighted areas.
3.7.15 References
"A Procedure for Measuring Extracted Chlorophyll a Free from the Errors Associated with
Chlorophyll b and Pheopigments", Turner Designs Method. 2006.
Arar, E.J. and Collins, G.B. "Method 445.0: In Vitro Determination of Chlorophyll a and
Pheophytin a in Marine and Freshwater Phytoplankton by Fluorescence", U.S. EPA
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methods for the determination of Chemical Substances in Marine and Estuarine
Environmental Samples. 1997.
Arar, E. J. "Method 446.0, rev 1.2: In Vitro Determination of Chlorophylls a, b, ci+ C2and
Pheopigments in Marine and Freshwater Algae by Visible Spectrophotometry". U.S.
Environmental Protection Agency, Office of Research and Development, National
Exposure Research Laboratory, Cincinnati, OH, 45268. 1997.
ASTM. American Society for Testing and Materials. Standard Specifications for Reagent Water.
D1193-77 (Reapproved 1983). Annual Book of ASTM Standards, Vol. 11.01. ASTM:
Philadelphia, Pennsylvania, 1991.
Welschmeyer, N.A. 1994. "Fluorometric Analysis of Chlorophyll a in the Presence of Chlorophyll
b and Pheopigments", Limnology and Oceanography, 39:1985-1992.
3.8 Microtiter Plate Enzyme-Linked Immuno-Sorbent Assay
(ALISA) for Microcystin
This section describes the process for performing Microtiter Plate ELISA of Microcystins using
the Abraxis Polyclonal ADDA kit at the USGS Organic Geochemistry Research Laboratory
(OGRL) in Lawrence, KS. Results are for water samples and concentrations are reported
between 0.1 Dug/Land 5.Oug/L without dilution. This method is suitable for water and algae
samples that have been lysed and/or filtered.
3.8.1 Definitions
Laboratory Duplicate- An identical sample; designated here by a letter extension, L, appended
to the log number.
ELISA- Enzyme-Linked Immunosorbent Assay.
IMN- Immunoassay for Microcystin analysis code.
Immunoassay Quality Assurance Sheet- Also called "the blue sheet." This is a sheet for
each project analyzed that has a list of samples analyzed that day. It is also used to track data
from the analysis, through data entry, through QA, and to the Project Manager's office to be
sent out.
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Data Entry Sheet- This is a sheet generated from Excel that provides the needed information
for the data entry person to be able to enter the concentrations into the database used to store,
retrieve, and print reports for all project data.
Project Quality Control Sample- Each project that is analyzed IMN analysis must have a QC
sample that contains a hit of microcystin from that project (so actual representative matrix of that
project). This QC sample is re-analyzed every time a sample from that project receives IMN
analysis.
Laboratory Spike- An identical sample; designated here by a letter extension, S, appended to
the log number. The final concentration will be 0.75ug/L of Microcystin-LR plus the ambient
sample concentration.
3.8.2 Safety Precautions
This SOP is to be used in conjunction with an approved Chemical Hygiene Plan. Also, consult
the Chemical Hygiene Plan for information on and use of all PPE.
The Stopping Solution consists of a weak acid. Do not allow it get on your clothes or yourself.
Wash the acid off immediately with copious amount of water.
3.8.3 Equipment and Supplies
Descriptions of commonly used pieces of equipment, their advantages and their limitations are
listed below.
• Samples: the Project Manager will provide a list of samples to be analyzed. They should be
located in Mac Shared to Lab.
• Project Quality Control Samples: These are designated by the person performing this SOP
as new projects come in. The samples are stored in the immunoassay refrigerator door for
easy access. The data from this sample is reviewed by the Quality Assurance checker.
• Microcystins Plate Kit (Abraxis): A commercially available kit that contains the antibody
coated microtiter plate and the reagents needed to perform the assay. A set of instructions
is included in the kit.
• Multichannel Pipette & Tips: An 8-channel pipette is used for this method. Familiarity of the
use of the multichannel pipette is necessary to achieve reliable results. Practice with water
if you have never used this before.
• Reagent Reservoirs (Costar Cat Number 4870): Plain plastic reservoir for reagents that
accommodate the use of a multi-channel pipette.
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• 100 |jl_ Pipette and Tips: For measuring and transferring standards, controls, and samples
into the antibody coated plates.
• 1000 uL Pipette and Tips: For diluting samples for reruns.
• Vortex Genie: For mixing dilutions.
• Adhesive Sealing Film (Parafilm) for Micro Plates (such as Rainin, non-sterile, Cat. No. 96-
SP-100): Used to cover plates during incubation.
• Orbital Shaker Table (such as American Shaker Table V, Model R4140): To be used for
mixing microtiter plates during incubations.
• Permanent Marker (Sharpie Fine Point): For labeling samples, bottles, plates and covers.
• Timer: For measuring incubation times.
• Distilled Deionized Water: For diluting samples.
• Paper Towels: For blotting the microtiter plates dry after washing.
• Plate Reader (such as Metertech, Model M965 AccuReader): Complete with Metertech PC
Mate software for operation of machine. This machine reads the microtiter plates.
• Computer: To operate plate reader and record data.
• Printer: To print the results.
• Immunoassay Quality Assurance Sheet (blue sheet): Found on ResLab under
Immunoassay Data. This is used by the QA checker and will be written on when results are
printed off.
• Data Entry Sheet: Found on ResLab under Immunoassay Data. This is where results are
input for the data entry person.
• Three Ring Notebook: For storage of data, found in room 313 on bookshelf.
• Test tubes: For dilutions, if needed.
• Log-in Records Binder: This contains the hard copies of log-in information used for checking
the accuracy of log-in against the labels on the sample bottles. This binder is stored in the
Project Manager's office (room 313) or in the lab by the plate reader.
3.8.4 Procedures
Note: Deviations from SOPs must be recorded in an appropriate instrument or work log.
Allow Immunoassay kits and samples to warm to room temperature before analyzing (about one
hour). Make certain there is enough of all the reagents to complete the number of analyses
before beginning, if not allow another kit to warm.
Preparation following the manufacturer's instructions:
1. Samples will undergo a total of three freeze-thaw cycles. For the first cycle, thaw (at room
temperature) the 250ml glass ambers bottles the samples were collected in. Aliquoted
approximately 10ml of each well-mixed sample into new, labeled 20ml glass scintillation
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vials, one per sample. Placed the 20ml scintillation vials in a freezer to complete the two
additional freeze-thaw cycles, for a total of three freeze-thaw cycles. All thaw cycles should
be completed at room temperature. After the last frze-thaw cycle, filter approximately 10 ml
of each sample through a new, syringe filter (0.45um) into a new, labeled 20ml glass
scintillation vial. (Norm-ject syringes and Whatman Glass Fiber syringe filters (25mm, GF
0.45um filter, catalog #6894-2504) or other similar alternative are acceptable.) One new
syringe and filter should be used per sample.
2. All immunoassays are categorized by project code and then numerical order; therefore,
separate the samples by project code first and include a QC sample for each project code.
For each set of 10 samples, the first and fifth samples are duplicates. One spiked sample
should be included for every regular sample.
3. On the ResLab, prepare the microcystin data entry sheet by entering the QC samples and
samples, date analyzed, project code, etc.
4. The pipettor, tips, sealing film, pens, test tubes, and reagent reservoirs are located in the
immunoassay drawers.
5. Using the PC Mate software, enter into the template, the location of the standards, controls,
and samples on the Microtiter Plate. The plate can be made up of rows of strips of 8 wells
that will snap into a blank frame. Label each row with a pen.
6. Analyze all five standards (0.00, 0.15, 0.40, 1.00, and 5.00) in duplicate. Space the
standards at the beginning.
7. The PC Mate software is on the immunoassay computer and is used for controlling the
microtiter plate reader and for calculating results. There is a manual for this software, and it
should be read before using the program.
8. Prepare the appropriate template and print it for reference for loading the standards and
samples. At this time enter plate name, file name, operator, etc. and save all the information.
File names include the date the assay was performed and includes the letter of the assay.
For example, since the analysis code for Microcystin is IMN, we use the N from IMN at the
end of the file name. Before the N, the seq name order should be year, month, and then
day. If the day was 1/20/05, the file name would be 05JAN20N.
9. Turn on the plate reader using the switch on the back to allow it to warm for at least 30
minutes. The plate reader may need to be turned on before the computer boots up so that
the computer can access and control the plate reader.
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Assay Procedure (There are also manufacturer instructions in the kit contents. All
procedures are in agreement with the manufacturer's instructions.):
10. Prepare spiked samples as called for on run sheet by adding 15 uL of a 25 ug/L microcystin-
LR standard solution to 500 uL of sample in a labeled LC vial. Cap and vortex. (Note on 25
ug/L microcystin-LR standard solution: This solution does not need to be made fresh daily.
Stock m microcystin-LR standard can be made by dilution in LC/MS grade methanol. Be
sure to check purity by UV-VIS extinction coefficient or LC/MS/MS. It is acceptable to make
batches from the stock in glass 2 mL LC vials and freeze them half-full. The final methanol
content is 5% or less in the 25 ug/L MCLR standard used for ELISA.)
11. Using the 100 uL pipette, add 50ul of the standards, controls, samples and spiked samples
(prepared in 6.3.1) to the appropriate wells in the plate as indicated by the IMN run sheet.
12. Immediately add 50uL of the pink antibody solution to each well using the multi-channel
pipette and a reagent reservoir.
13. Place Sealing Parafilm over the wells.
14. Place the plate on the orbital shaker table. Protect tray from light, and set the speed for 180
rpm and the timer for an hour and a half.
15. Carefully remove the Parafilm.
16. Empty the plate into the sink, pat dry with a stack of paper towels, and then wash the wells
of the plate 3 times with 250ul of washing solution using the multi-channel pipette. Each
time you add the washing solution, let the washing solution set about 45 seconds before
emptying it into the sink and using the paper towels as before.
17. Add 100ul of enzyme conjugate solution to all wells using the multi-channel pipettor.
18. Cover the wells with Parafilm and place on the orbital shaker table in a dark corner at 180-
rpm for 30 minutes.
19. After 30 minutes, rinse the wells 3 times again with 250ul of washing solution.
20. Add 100ul of substrate solution to the wells using the multi-channel pipette and reagent
reservoir, and incubate on the orbital shaker at 180 rpm for 25 minutes. Cover with Parafilm
and protect from light. This will make the contents have a blue hue.
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21. Uncover and add 50ul of stopping solution to the wells. This will turn the contents a bright
yellow color. After you have added the stopping solution, the plate must be read within 15
minutes using the plate reader.
Reading the Plate
22. Change the settings in the PC Mate program if needed to agree with the manufacturer's
instructions. The current settings for the Microcystin Assay are:
• Wavelength 450nm
• Curve is: 4-parameter fit
• Display: Analyzed
• Endpoint: L1, Auto mix: on, cal:on, disk:on, print:OFF
Reporting Data
23. The software calculates the values of the samples from the Calibration Curve, and
averages the two results. Print this report out.
24. Reporting limits for this procedure range between 0.10 ug/L to 5.0 ug/L. If a dilution is
performed, higher concentrations can be reported. Non-detections are reported as "<0.10
ug/L".
25. Prepare the Data Entry Sheet if you have not already.
26. There is an Excel workbook in ResLab labeled Immunoassay Data. Copy a previous sheet
and type today's information into it, including Lab ID, project code, concentrations to report
(get this information from the raw data print out), analysis date, any remarks, your initials as
the tech ID, etc. The entire plate's samples go on a single Date Entry Sheet (i.e.: it can
have multiple projects on it).
27. An example of the Data Entry Sheet is shown in Attachment C.
28. When typing in the concentration of the sample, you will find that there are three values on
the strip. Use the value that says the mean concentration, the last value of the three. This
will be input into your data entry sheet. The strip will show if it is "HI" for out of range values.
29. Samples needing dilutions because of "HI" concentrations do need QA's approval and can
be redone in the next run. The same is true for any other redo, such as if the duplicates did
not agree, you ran out of stop solution, or there was a bad standard curve.
30. For non-detects (flagged as "nd") list as less than the reporting limit ("<0.10").
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31. You should also enter "DO NOT ENTER" for samples that had high concentrations, needed
dilutions, or ran out of reagent, as well as for QC samples. QC values go in a separate data
base.
32. To determine if the control is correct, type in the found value in the analyzed column (in blue
font at the bottom of the page). It will calculate a percent difference which must be within
20% of the control concentration.
33. Print out the sheet and attach it to the blue sheet.
34. Print out the computer information and attach it to the blue sheet (include all calibration
curves and values).
35. Complete the Immunoassay Quality Assurance Sheet (i.e. the blue sheet).
36. This is in the same Excel workbook as the data entry sheet. Copy over the previous sheet
with new information. Prepare a new blue sheet for each project analyzed on this plate. You
must type in the project code, analysis code (IMN), plate name, analysis date, and the lab
ID's. Print the sheet(s) out on blue paper, which is kept in drawer LciC11&12 with the other
colored paper.
37. Do a bottle check (a verification of the log-in accuracy) on the samples analyzed.
38. Obtain the 3-ring binder of Log-In records from the Project Manager's office (Rm 313).
39. Using the Lab ID's (MM#'s) to guide you, compare the information printed in the binder to
what is on the bottle and look for discrepancies. Check the project code, site name, station
ID, date collected, and time collected.
40. Place a check mark (S) next to the Lab ID in the binder that corresponds to the bottle you
just verified to signify that the log-in information has been checked against the bottle.
41. If you find a mistake or discrepancy, write in the correction in the binder, initial your
correction(s) in the binder, immediately report it to the Project Manager, and see that it gets
corrected in the project's database computer.
42. Initial and date the Immunoassay Quality Assurance Sheet (blue sheet) to indicate that
sample labels have been verified against log-in and corrections made.
43. An example of the Immunoassay Quality Assurance Sheet is shown in Attachment B.
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44. You now have a data set consisting of the raw data (plate template, concentrations, and
standard curve) printed from SoftMax, the Data Entry Sheet, and the Immunoassay Quality
Assurance Sheet. Paperclip these together with the Immunoassay Quality Assurance Sheet
on top and turn it in to the Data Entry box for IM analyses (Rm 309).
45. Dispose of solution in plates in a lab sink. Rinse the plate and sink with water to dilute the
weak acid present.
46. All data from the analysis is put in the IMN folder, which is found in the Project Manager's
office. That report includes the Curve that is printed out from the computer, the data sheet,
and the template for the standards and the samples. Staple this information together.
Dilutions if needed are prepared as follows (using clean disposable plastic tubes):
47. 1. 1:10 dilution
• Pipette 100ul from the sample in to the tube.
• Add 900 ul of distilled water to sample above. (Note: Dilutions may also be
made using the kit's dilutenet rather than distilled water.)
• Mix by Vortexing.
• Multiply final concentration by 10.
48. 1:100 dilution.
• Pipette 10ul from the sample.
• Add 990ul of distilled water to sample. (Note: Dilutions may also be made using
the kit's dilutenet rather than distilled water.)
• Mix by Vortexing.
• Multiply final concentration by 100.
49. Other dilutions can be calculated if needed.
3.8.5 Quality Control
The Quality Assurance Officer or designee will evaluate overall data quality and QC compliance.
In the event data is not in compliance, the problem(s) will be identified and samples will be
reanalyzed as appropriate after corrective action taken.
The standard curve should have a correlation coefficient of 0.99 (as suggested by ELISA kit
manufacturer).
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The absorbency of the blank must be >1.400 (as suggested by ELISA kit manufacturer).
The Check Standard supplied with the ELISA kit should be analyzed a minimum of two times in
each run, once at the beginning and once at the end. This helps ensure the plate was prepared
in the proper time frame. Values should be +/- 20% (28.3% relative standard deviation (RSD))
of expected value.
Laboratory duplicates should have a percent Relative Standard Deviation (% RSD) of 28.3
percent or less when compared to each other (as suggested by ELISA kit manufacturer).
If laboratory duplicates are outside of this range, then they should be reanalyzed in the next run.
Quality control samples are available for each project. Criteria for acceptance of measured
values are +/- 20% of average. These samples are analyzed everytime samples from the same
project code are analyzed.
A designated archived project sample (Quality Control Project Samples) is reanalyzed with
every run set that is analyzed. Control charts are maintained for these samples. A running
historical average is maintained of the concentration from each run. The concentration of the
QC sample for each successive run has to be ± 20 percent of that average to be acceptable.
3.8.6 References
USGS. 2010. Standard Operating Procedure: Microtiter Plate Enzyme-Linked Immuno-Sorbent
Assay for Microcystin. OGRL-SOP-5630.
3.9 Taxonomic Standards Protocols
The taxonomic nomenclature for the NWCA will follow previous lists that have been developed
for the 2008 National Lakes Assessment, the National Rivers and Streams Assessment, and the
USGS National Water-Quality Assessment (NAWQA) Program list (Appendix B). These three
lists have not been reconciled but the names used are sufficiently consistent. These lists include
accepted names for all distinct taxa for both known and unknown species. The laboratory is
responsible for reconciling all identifications with these taxa lists. Voucher images are reviewed
regularly to ensure accuracy and consistency among taxonomists.
If a species of algae cannot be conclusively determined, then several strategies can be used to
characterize what is observed.
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1. If a taxon is observed repeatedly and can be consistently distinguished from other diatoms,
then it is given a Counter Taxon Name designation in the following format:
Unknown Species
a. Genus name
b. "X" is a unique sequence number for unknown species in that genus
c. NWCA 11 is the unique project identifier
d. "YYY" are the initials of the taxonomist who first made the identification
e. Example species name:
i. Nitzschia sp. X NWCA11 YYY
ii. Nitzschia sp. 1 NWCA11 EAR
iii. Nitzschia sp. 2 NWCA11 EAR
iv. Acanthosphaera sp. 1 NWCA11 EAR
v. Cyanobacterium sp. 1 NWCA11 EAR
The Counter Taxon Name designation can also be used at higher levels of taxonomy in the
following format:
Unknown Genus
a. "Unknown" identifies the species as having an unknown genus
b. Division name (i.e., Rhodophyta or Chlorophyta)
c. Brief description of algae to help taxonomists distinguish between unknowns
d. "X" is a unique sequence number of unknown species of that division
e. NWCA11 is the unique project identifier
f. "YYY" are the initials of the taxonomist who first made the identification
g. Example species name:
i. Unknown Chlorophyta coccoid sp. X NWCA11 YYY
ii. Unknown Chlorophyta coccoid sp. 1 NWCA11 EAR
iii. Unknown Rhodophyta flagellate sp. 1 NWCA11 EAR
2. If a taxon looks similar to another taxon, but not sufficiently to be identified as that taxon,
then the "cf" designation is used in the following format:
a. Genus name
b. "cf" designates the taxon as not sufficiently identified
c. Species name
d. Last name of the taxonomist to first use the designation in capital letters
e. Species authority
f. Example species name:
i. Chroococcus cf. minimus CIUGULEA (Keissler) Lemmermann
3. If a taxon was only observed in one sample and can be distinguished from other algae in the
sample, but it is not tracked in other samples or cannot be related to another taxon, then it is
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assigned as "sp. X ?", where X is a unique sequence number. This designation can be used
with genus or higher levels of taxonomy, e.g. Unknown Cyanophyte sp. 1?
4. If a specimen cannot be identified to species level or distinguished clearly from other
unknowns in a sample, then the designation "sp." or "spp." is used. The designation "sp." is
used when there is likely only one taxon of undistinguishable cells in the sample (e.g.,
Dictyosphaerium sp.). The designation "spp." is used when there is likely more than one
taxon of undistinguishable cells in the sample (e.g., Dictyosphaerium spp.). These
designations can also be used at higher levels of taxonomy, but it is important to include cell
shapes or growth forms so that taxonomists can distinguish between unknowns, e.g.
Unknown Chlorophyte coccoid (>10u) GDL or Unknown Chrysophyte flagellate.
Girdle views of diatoms are often difficult to identify to species level. However, taxonomists for
the NWCA samples should identify diatoms in girdle view to species and variety level based on
their knowledge of the taxon in valve view and their ability to understand the three dimensional
morphology of a specimen.
3.9.1 Images
Taking pictures, illustrating taxa, and maintaining an image database is important for consistent
identification of taxa by individuals and groups of taxonomists. The NWCA protocol requires
good quality digital images to be supplied for all taxa that represent 5% or greater abundance of
any one sample. Good quality means that the taxon can be identified under the microsope from
that digital image. Unknown taxa also must be imaged and shared with other taxonomists.
Images of all unknown taxa given NWCA species designations shall be provided to document
these taxa. Images shall be shared among taxonomists so all taxa, named species, and NWCA
unknowns, can be consistently identified in samples by more than one taxonomist. These
images are critical for increasing consistency in taxonomy, i.e. ensuring that the same name is
used for commonly occurring 'unknowns' throughout a project.
Images should be taken at a focal depth at which most key diagnostic features are
distinguishable and clear. Flat specimens are preferred over tilted specimens. They should be
taken at the magnification that the samples were counted. Images should be cropped to include
only one taxon. The image should include a scale bar or measurements to indicate the size of
the specimen. Consistent labeling of images is helpful when maintaining an image database.
The naming convention used for images in the NWCA image database shall be as follows:
Taxa name_magnification (if applicable)_sample ID.xxx
e.g. Homoeothrixjanthina_40x1.5x_NWCA0001 Jpg
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3.9.2 Table of References
Documentation of the concepts used for all taxa is important. If it is not practical to take a good
digital image of certain taxa, the taxonomist should provide an image from a reference that best
represents the morphological concept of the taxon as used in the study. The taxonomist is
responsible for developing a NWCA Reference Table, which lists all of the NWCA Taxa Names,
references used to identify the taxa, and an image from a plate in the literature or collection of
digital images. Literature and image references will be provided for all taxa with NWCA Taxa
Names at the species or variety level and all NWCA unknown taxa that are category 1
unknowns, e.g. "Ankistrodesmus sp. 1 NWCA JAM." Each species identified at the species or
variety level with a species name or identified as a distinct unknown is incorporated in the
reference table.
3.9.3 Taxonomic Harmonization
Successful taxonomic harmonization among taxonomists is essential for accurate data analysis.
Inconsistent identifications of algae introduce error in ecological analyses, so reducing this
source of error is important. Harmonization of identifications among taxonomists is particularly
important in large projects in which more than one taxonomist identified algae. The process of
harmonization can be accomplished without limiting the taxonomic explorations of taxonomists.
To accomplish both goals, the lab develops harmonization tables in which names used in
identification by each taxonomist are related to names that other taxonomists used in the
project.
Taxonomists will participate in conference calls as needed or at least once a month to
harmonize taxa for known diatoms and soft algae. Unknown species are also reconciled to a
standard naming convention during this time. During these conference calls, taxonomists share
species images for 1) all unknown taxa given Counter Taxon Names, and 2) known species that
occupy 5% or great of either count.
During the harmonization process, names assigned by taxonomists during counting, referred to
as Counter Taxon Names, are assigned NWCA Taxon Names based on taxonomist agreement.
For known taxa the Counter Taxon Name and the NWCA Taxon Name may be the same.
However, in some instances a NWCA Taxon Name may correspond to one or more Counter
Taxon Names, (i.e., multiple taxonomists recognize the same unknown taxa). For unknown taxa
name harmonization use the following standard naming convention:
NWCA Taxon naming convention for unknown species:
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a. Genus name
b. "X" is a unique sequence number of unknown species of the unknown genus; note this
number sequence restarts during the taxonomic harmonization process. For example, if
taxonomist A's first Nitzschia species and taxonomist B's twelfth Nitzschia species are
the same, then they would become the first harmonized Nitzschia species (i.e, Nitzschia
sp. 1 NWCA11 EAR and Nitzschia sp. 12 NWCA11 MES become Nitzschia sp. 1
NWCA11 LAB).
c. NWCA11 is the unique project identifier
d. Name of lab identifies that the unknown species has been reconciled during the inter lab
taxonomic harmonization process
NWCA taxa that are likely too similar for multiple taxonomists to name consistently are assigned
a Lump Level 1 Name. Lumping similar taxa will reduce taxonomic variability and reduce the
risk of error in the ecological analyses. This process is repeated to create a Lump Level 2
Name, which represents the coarsest level of species level taxonomy recommended for use in
ecological analyses. With each level of harmonization, reasons for lumping at successively
higher levels should be provided.
The product of the harmonization process is a Taxonomic Harmonization Table that provides
the following information:
1. Counter Taxon Name
2. Reason for lumping Counter Taxon Name to an Lab Taxon Name, if it was lumped
3. NWCA Taxon Name
4. Reason for lumping NWCA Taxon Name to a Lump Level 1 Name, if it was lumped
5. Lump Level 1 Name
6. Reason for lumping Lump Level 1 Name to a Lump Level 2 Name, if it was lumped
7. Lump Level 2 Name
8. Comment
3.9.4 Final Products
The lab is responsible for sending the following data files to the Information Management
Coordinator at NARSDataSubmission(S)epa.gov for the purpose of quality assurance: count
tables in comma delimited text files or Access database tables, taxonomist reports, and
harmonization (translation) tables. A written report detailing the methods used and discussing
problems encountered must be included as an appendix to this report. In addition to these data
files, the lab will send the reconciled and harmonized species list as a final product. All products
should be provided in the same format as the National Rivers and Streams Assessment
products.
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3.10 Pertinent QA/QC Procedures
The QA/QC procedures for algae include taxonomic harmonization and re-counts of 10% of
samples collected. These procedures should occur continuously throughout the project to
correct for inaccuracies and ensure the final products contain high quality data. For re-counts of
samples, analysts will swap vials of material and hard mounted slides for diatoms and recount
following sample protocols. Re-counts will be performed by an inter-lab exchange. EPA will
inform the laboratories which random samples will be re-counted. The samples must then be
sent from the original laboratory to another laboratory. The QC taxonomist should complete
another copy of the Taxonomic Bench Sheet for each sample. Each bench sheet should be
labeled with the term "QC Re-ID." As each bench sheet is completed, it should be emailed to
the Information Management Coordinator at NARSDataSubmission(S)epa.gov.
The lab will compare the taxonomic results generated by the primary and QC taxonomists for
each sample based on both the raw data and the appropriate metrics (i.e., taxa identified with
similar autecologies). EPA will then calculate percent similarity (Algae Chapter of QAPP). It is
expected that the soft algae counts should have a similarity of >50% and the diatom counts
should have a similarity of >70%. If not, the reasons for the discrepancies between taxonomists
should be discussed. Results less than these values will be investigated and logged for
indication of error patterns or trends.
A report or technical memorandum will be prepared by the QC Taxonomist. This document will
quantify both aspects of taxonomic precision, assess data acceptability, highlight taxonomic
problem areas, and provide recommendations for improving precision. This report will be
submitted to the Information Management Coordinator, with copies sent to the primary and QC
taxonomists and another copy maintained in the project file. Significant differences may result in
the re-identification of samples by the primary taxonomist and a second QC check by the
secondary taxonomist. All samples must be stored at the laboratory until the project officer
notifies the lab.
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4. VEGETATION
4.1 Introduction to Indicator
Collectively, wetland plant species 1) represent diverse adaptations, ecological tolerances, and
life history strategies, and 2) effectively integrate environmental conditions, species interactions,
and human-caused disturbance. Data describing species composition and abundance and
vegetation structure are powerful, robust, and relatively easy to gather (USEPA, 2002).
Categories of data to be collected include the following:
1. Species composition and abundance
2. Alien species
3. Floristic quality
4. Guild composition
5. Community composition
6. Vegetation structure and productivity
For more detailed information on the utility of vegetation for assessing wetland condition please
see Ecological Indicators for the 2011 National Wetland Condition Assessment (in preparation).
4.2 Receiving Voucher Samples
Processing and managing plant samples or identifying unknown species in the field is often very
difficult and time consuming. Accordingly, it can be logistically infeasible to complete, along with
vegetation sampling tasks, in a single day. Thus, it is extremely important that the Vegetation
Team is prepared to ship samples to a herbarium or bring them back to the office to complete
this task. Plant samples will arrive at the herbarium dried and pressed in shipping boxes.
4.2.1 Definitions
For the NWCA, a voucher sample is a pressed and dried plant sample, ideally comprised of
leaves, stems, flowers, fruits and roots. An integral component of each voucher sample is
written data describing the location, date of collection, habitat, plant habit, characteristic
features, and other information. Vouchers provide physical evidence that confirms the presence
of plant species at specific locations. Other uses for vouchers include 1) identifying species
encountered and not immediately recognized by a botanist or other field workers during
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ecological studies, 2) documenting the occurrence of native rare plants, or 3) cataloging the
distribution of invasive alien plants.
For all NWCA field work, whenever the identity of a species cannot be confirmed in the field, a
sample is collected (see Vegetation Chapter of Field Operations Manual (FOM)) for later
identification in the office or lab. All unknown species in each Vegetation Plot (five 100-m2 Veg
Plots/AA) are collected. If an unknown species is collected at a previous site, it is collected at
subsequent sites, until the Botanist/Ecologist learns the identity of the species and can reliably
sight recognize it in the field. This is particularly important for species in difficult wetland genera
and families, such as those that include sedges, rushes, grasses, and submerged aquatic
vegetation. The Botanist/Ecologist will ship unknown samples to the Herbarium for identification
(Vegetation Chapter of FOM).
In addition, one known species is randomly collected from each Vegetation Plot as quality
assurance voucher samples (Vegetation Chapter of FOM). Collecting voucher samples of
known species both provides a quality assurance check on species identity data, and a
permanent record of the occurrence of a particular species at a given location. The field
Botanist/Ecologist will ship known samples collected for quality assurance to the QA Herbarium.
One of the primary goals of the national survey is to build state and tribal capacity to implement
wetland monitoring and assessment programs; therefore, EPA encourages states to participate
in the vegetation portion of the lab manual wherever possible. For the purposes of this manual a
Herbarium represents the person identifying and processing unknown samples. This could be a
field Botanist/Ecologist, state identified herbarium, EPA identified regional herbarium, or
National EPA Contractor. The Herbarium is responsible for ensuring all plant identification and
processing tasks outlined in this manual are completed. In some cases this may require the
Herbarium to identify partners to assist with the work.
Voucher samples collected for quality assurance will be sent to the designated QA Herbarium.
A QA Herbarium is a certified botanist, state or EPA identified herbarium that agrees to use the
NWCA prescribed methods, as described in Section 4.7 of this chapter, to ensure that all QA
vouchers receive the same level of taxonomic precision. If the Herbarium and QA Herbarium
are the same institution, it is important that all Quality Assurance activities are completed by
another experienced taxonomist who did not participate in the identification of unknown species.
The QA Herbarium will blindly re-identify all species to ensure that the identifications are
independent.
See figure 4-1 for potential options for plant samples collected in each Vegetation Plot.
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la)- Field Crew
Botanist/Ecologist, or Other
State Botanist or Herbarium
selected by State to identify
unknowns
2a) - Quality Assurance Species
(5 randomly selected species of known
identity for QA verification, 1
species/Veg Plot)
Ib) - EcoAnalysts Plant Lab
3} - Independent Botanist
*For specimens from 2a - not the person
who collected the QA specimens in the field
•For specimens from 2b: - not the person
who identified these species
2b) -10% of species identified in steps la or
Ib, randomly selected for QA verification
Figure 4-1: Potential options for plant vouchers. NWCA definitions for Herbarium and QA
Herbarium are provided in section 4.2.1.
4.2.2 Tracking information
In the field, each voucher sample collected is assigned a set of tracking information, which is
recorded on the Plant Sample Tracking Form (Figure 4-2a and 4-2b). At the end of the sampling
week the vegetation team will remove the samples and newspaper sleeves from the press,
ensuring they retain the Plant Sample Label Form (Figure 4-4), and ship them in a sturdy box to
the Herbarium (Vegetation Chapter of FOM). If a sample listed on the tracking form is not part of
the shipment, or a sample arrives at the lab without the proper label, contact the Information
Management Coordinator (541-754-4663) as soon as possible.
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A FORM T-2: NWCA 2011
Sent By: Sender Phone:
UNKNOWN PLANT SAMPLE TRACKING R»I-«I A
State Where Sites are Located:
Shaped Using: O FedEx O Other:
Airbill/Tracking Number
Retained for Identification By:
OR
Phone Number:
Date Shipped: / / 2 0 1 1 Date: / / 2 0 1 1
Instructions:
1. Complete all header and shipping information above.
2. Fill out the body of the form with Site ID, Visit Number and date collected. List the plant samples for unknown species b, ' jllection
number (listed on Form V-2, and on plant sample tags and labels! or by 'anges ot cD:lection numbers (e.g., I-3, 5-L S-] '., 13, 15-
19(. If needed, more than one row can be used for a site. Be sure to record the Site ID for each line.
3. For shipped plant samples, place photocopy of this form in shipping package. For retained plant sample s, * 'ad f py of this form in
folder or box in which they are stored.
4. Send original form to the Information Management Team as soon as the plant samples are shipped to he designated Lab or
transferred for storage by the State Team for later identification.
2011 Collection
S ile ID Vis it * Date I MM.'D D) Plant Sampl e Colection Numbers
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
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NWCA11-
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MWCA11
NWCA11-
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Chain of Custody Tracing Assistance:
Q EcoAnalyst Plant Lab:
Address info unknown
O State Plant Lab: MarV5 Cappaert
ph: 541-754-4467
Will be prefilled based
on info provided Michelle Gover
ph: 541-7&M793
Sample Tracking Contact Info:
FAX: 541-754J637 VOICE MESSAGE CENTER: 541-754-4663 EMAIL: sampletrackingQepa.gov 58 64386613
NWCA-raatlng-Uh>K Plant Sample
Figure 4-2a: Unknown Plant Sample Tracking Form
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• FORM T-3: NWCA 201 1 QA PLANT SAMPLE TRACKING >*""»i Jfc
Sent By: Sender Phone:
Shaped Using: O FedEx O Other:
Airb ll.'Track ng Number
0
Date Shipped: / / 2 0 1 1
State Where Sites are Located:
Retained for Identification By:
R
Phone Number:
Date: / / 2 0 1 1
Instructions:
1. Complete all header and shipping Information above.
2. Fill out the body of the form with Site ID, Vls.lt Number and date collected. List the QA plant samples for species identify I . the
field and/or the QA samples far plant species identified In the Lab 'by collection number (listed on Form v-2 and on plan* sample
tags and labels). Collection numbers can be listed Individually (e.g., 4, 6, 1Q\ oc as ranges (7-12) tf appropriate. If -.eeded, more
than one row can be used for a site.
3. For shipped plant samples, place photocopy of this form in shipping package. For retained plant samp' _s. pk.-ei.jpy of this
form In folder or box In which they are stored.
4. Send original form to the Information Management Team assaoa as the plant samples are shipped to t, « designated Lab or
transferred far storage by the State Team for late- identification to allow proper tracing of plant • n|-l..v..
2011 Collection
Site ID Visit* Date|MM,'DD) Plant Sample Coleciion Numbers
NWCA11-
NWCA11-
NWCA11-
NWCA11
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
NWCA11-
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CM
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Chain of Custody
O EeoAnalyst Plant Lab:
Address info unknown
Q State Plant Lab:
Will be prefilled based
on info provided
Trackiryg Ass-stance:
M:-'y= Cappaert
ph: 541-7544487
Michelle Cover
ph: 541-7644793
Sample Tracking Contact Info:
FAX: 541-754-4S37 VOICE MESSAGE CENTER: 541-754-4663 EMAIL: sampletrackingttepa.gov
11/292010 NWCATtaOilng-QA Rant Sample
073146971E
Figure 4-2b: QA Plant Sample Tracking Form
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4.3 Supplies and Equipment for Sample Handling
• Plant dryer
• Dissecting microscope
• Dissecting tools (e.g., single edge razor blades, forceps, dissecting needles)
• Regional floras and plant lists
• USDA PLANTS taxonomic standard (http://plants.usda.gov/java/)
• Plant nomenclatural forms
• Plant sample tracking forms
• Plant sample folders
• Storage cabinet or sealable plastic boxes for storing dried plant samples prior to
identification
• OPTIONAL: Freezer or naphthalene for killing pests on dried plant material
• OPTIONAL: Mounting materials (herbarium sheets, mounting glue, forceps, weights for
holding samples with wet glue to the herbarium sheets, etc)
• OPTIONAL: Herbarium sample labels
4.4 Handling Vegetation Samples
Plant samples may arrive at the Herbarium in several conditions: 1) as dried, pressed samples,
or 2) pressed but still wet plant material enclosed in a plant press.
1. If samples are pressed and dried, proceed to Section 4.4.3 (Treat samples for detritivores,
molds, and pests), then to Section 4.5 (Identifying Plant Samples).
2. If samples arrive in a press, but are still wet they should be placed on a plant dryer to
complete drying, and then be treated for pests. This would likely occur if the
Botanist/Ecologist is acting as the Herbarium.
4.4.1 Plant Sample ID Tag and Label Form
Every sample will arrive at the Herbarium with a Plant Specimen Label Form. This form includes
diagnostic information for known and unknown species collected. Voucher samples are
considered incomplete without this information. An example of the Plant Sample ID Tag Form is
provided in Figure 4-3 and an example of the Plant Sample Label Form is provided in Figure 4-
4. The Plant Sample ID Tag will be affixed to the outside of the newsprint in which each
specimen is pressed. A duplicate tag will be affixed to the Plant Specimen Label. If a sample
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does not have any of the following information, contact the Information Management
Coordinator (541-754-4663) as soon as possible:
Plant Sample ID Tag Information
• Plant Sample ID Number: NWCA Site Number-Plant collection number. Plant collection
numbers for samples are ordered chronologically for each site beginning with one. For
example, the sample number for the 5th species collected at NWCA11-Point 0762 would
be NWCA11-0762-5. Voucher samples collected for QA purposes will have the QA
bubble filled.
• Collection Date: Date is numerical: month, day, year, e.g. 06/14/2011.
Plant Specimen Label Information
• Plant Sample ID Number: NWCA Site Number-Plant collection number. Plant collection
numbers for samples are ordered chronologically for each site beginning with one. For
example, the sample number for the 5th species collected at NWCA11-Point 0762 would
be NWCA11-0762-5. Voucher samples collected for QA purposes will have the QA
bubble filled.
• Collection Date: Date is numerical: month, day, year, e.g. 06/14/2011.
• Pseudonym for Unknown Species: Descriptive name used on data forms (e.g., Carex
sp. 1). QA specimen will not have species name information on the label.
• Collector(s) name: List the first name, middle initial and surname of the person or
persons who collected the sample.
• Habitat: The type of plant community or setting where the plant is growing, (e.g., such as
wetland type (Cowardin, HGM, NVC), wetland community type (forested wetland,
emergent marsh, wet prairie, mountain bog, etc.), anthropogenic disturbances (urban
setting type), and, other plants growing in association (associated species information
would be available from the plot).
• Plant habit/description: Describe key features of the plant such as growth form (tree,
shrub, vine, herb), approximate height, longevity (annual, biennial, perennial), clonal,
rhizomatous, tussock-forming, etc. List any characteristics of the plant which may be lost
upon drying, such as flower/fruit color, fragrance, and leaf orientation.
• Abundance of Plant: Indicates whether the species is dominant, common, sparse or
uncommon at the site, and whether based on canopy cover it is a dominant, moderate
cover, or low cover species.
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Plant Sample 10 Number:
(Sites) (Collections)
Date: Q2Jtfj2Qll Visit #: •! O2
• - fill in circle if QA specimen
Figure 4-3: Example Plant Sample ID Tag
PLANT SPECIMEN LABEL (enclose in newsprint with sample)
For Unknown species, record pseudonym from Form V-2:
Collector^) Name(s):
L.
Plant Sample ID Number:
(Site«)
(Collection 8)
Date: (??/ OH/2O11 Visit »:•! O2
O - fill in circle if QA specimen
Abundance of Plant (fill appropriate circle):
ODominant, •Common, OSparse, OUncommon
Habitat:
Op So/i_'WfT
&RS Ł Ł>
tJf-ft
Plant Habit:
M/
/Ł*!/ŁŁ
8 *Sf-L
Figure 4-4: Example Plant Specimen Label
4.4.2 Drying Samples
Plant samples arriving at the Herbarium may still be wet and in the plant press (this would likely
happen if the Botanist/Ecology is acting as the Herbarium). The pressed plants must be
thoroughly dried before removing them from the presses. As the samples dry they will loose
volume, so it is often necessary to periodically tighten the straps on the press to maintain
pressure on the samples and minimize shrinkage and wrinkling.
Low ambient humidity and good airflow around and through the presses is important for rapid
and thorough drying of plant material. Rapid drying over low heat promotes preservation of color
and morphology resulting in high quality samples. Dry air circulating through the press also may
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kill many insects and insect eggs, which may protect the samples from some insect damage.
These conditions are most easily obtained by placing full presses on an electric plant dryer that
provides steady bottom heat (95°F to 113°F), where plants usually dry in 12 to 48 hours.
However, presses placed in a warm dry place will be sufficient if a plant dryer is not available.
4.4.3 Treat samples for detritivores, molds, and pests
Dried plant material is highly susceptible to contamination by detritivores, molds, and pests that
can destroy herbaria collections. Therefore it is important to treat all incoming samples to kill
potential contaminates.
Herbaria should implement their standard pest procedures. A common method for sample
treatment is to freeze them (-20°C or below) for at least three days for loosely stacked samples
and seven days for tightly packed samples.
To protect the collection from infestation, plant samples should be stored in herbarium cabinets
or sealable plastic container when not in use; under no circumstances should samples be left
out in the herbarium room overnight. If samples are found that have been left out overnight or if
a cabinet/plastic container has been left open, all samples may need to be decontaminated
again.
4.5 Identification of Vegetation Samples
4.5.1 Taxonomic standard
The recognition and identification of particular classes of plants such as families, genera, and
species is a critical and difficult element of collecting accurate vegetation plot data. To
complicate matters, not all botanical authorities agree about which name to apply to a particular
plant species. The NWCA is using the taxonomic nomenclature of the USDA Plants Database
as the taxonomic standard. To effectively key plants and identify them in the field, however,
Field Crews may use local floras appropriate to each region or state (Appendix C). This means
numerous taxonomies will likely be applied across the 48 conterminous states comprising the
study area. The Herbarium will reconcile all species names that they identify to the standard
found in USDA Plants.
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4.5.2 Species Identifications for Voucher Samples
Once each plant specimen is identified, the taxonomist is responsible for reconciling the species
name with the USDA Plants standard. All identifications are recorded in an excel database. Use
the following steps to compare the final taxa list for each site to that of the USDA Plants website
(http://plants.usda.gov/).
4.6 Mounting and Storing Herbarium Sheets
Once the samples are dried, pressed, and identified, they are to be stored at the Herbarium for
at least five years. Vouchers should be kept in sealable plastic containers in a cool dry climate
and must be accessible to the EPA. However, the Herbarium is encouraged to incorporate the
NWCA vouchers into their permanent collections as desired. Vouchers from the national survey
mounted on herbarium sheets should be labeled to indicate that they were collected as part of
the NWCA. For an example of commonly used mounting and labeling methods see Appendix
D.
4.7 Quality Assurance
A subset of plant samples collected as unknowns and later identified by the Herbarium will need
to be verified by a QA taxonomist for additional Quality Assurance (QAPP section 5.1.8). The
Herbarium will randomly select 10% of the samples to be sent to the QA taxonomist, another
experienced taxonomist who did not participate in the original identifications.
Note: If the field botanist/ecologist is acting as the Herbarium then the QA Herbarium
would need to be another qualified botanist, or a state or EPA identified herbarium.
However, if a state or EPA identified herbarium is acting as the Herbarium then the QA
activities could be completed by a taxonomist within the same institution that has not
actively participated in the identification of the unknown species.
1. The QC taxonomist will perform re-identifications completing another copy of the Vegetation
Taxonomic Bench Sheet for each sample. Each bench sheet must be labeled with the term
"QC Re-ID." As each bench sheet is completed, it must be faxed to the Information
Management Coordinator.
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2. The project facilitator will compare the taxonomic results generated by the primary and QC
taxonomists for each sample and calculate percent taxonomic disagreement (PTD) as
measures of taxonomic precision (Stribling et al. 2003) as follows:
fompDOS
PTD =• -1 P 100
_ N
Equation 1
where compp0s is the number of agreements (positive comparisons) and N is the total
number of samples in the larger of the two counts.
3. Unless otherwise specified by project goals and objectives, the measurement quality
objective for enumerations will be a mean PTD less than or equal to 15, calculated from all
the samples in the 10% set sent to the QC taxonomist. Results greater than these values
will be investigated and logged for indication of error patterns or trends, but all values will
generally be considered acceptable for further analysis, unless the investigation reveals
significant problems.
4. Corrective action will include determining problem areas (taxa) and consistent
disagreements. Problems will be addressed through taxonomist interactions.
Disagreements resulting from identification to a specific taxonomic level, creating the
possibility to double-count "unique" or "distinct" taxa will also be rectified through corrective
actions.
5. The project facilitator will prepare a report or technical memorandum. This document will
quantify both aspects of taxonomic precision, assess data acceptability, highlight taxonomic
problem areas, and provide recommendations for improving precision. This report will be
submitted to the project manager, with copies sent to the primary and QC taxonomists and
another copy maintained in the project file.
4.8 References
Kansas State University. 2006. Integrated Pest Management Plan, http://www.k-
state.edu/herbarium/pests.html. Accessed November 17, 2009.
Lee, M. T., R. K. Peet, S. D. Roberts, and T. R. Wentworth. 2008. CVS-EEP protocol for
recording vegetation: All levels of plot sampling. Version 2008. The Carolina Vegetation
Survey (CVS, http://cvs.bio.unc.edu) and the North Carolina Ecosystem Enhancement
Program (EEP, http://www.nceep.net).
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National Wetlands Condition Assessment
Laboratory Methods Manual Page 69 of 256
Mack, J. J. 2007. Integrated Wetland Assessment Program. Part 9: Field Manual for the
Vegetation Index of Biotic Integrity for Wetlands v. 1.4. Ohio EPA Technical Report
WET/2004-9. Ohio Environmental Protection Agency, Wetland Ecology Group, Division
of Surface Water, Columbus, Ohio.
Magee, T. K., S. E. Gwin, R. G. Gibson, C. C. Holland, J. E. Honea, P. W. Shaffer, J. C.
Sifneos, and M. E. Kentula. 1993. Research Plan and Methods manual for the Oregon
Wetlands Study. EPA/600/R-93/072. Environmental Protection Agency, Environmental
Research Laboratory, Corvallis, Oregon.
Stribling, J.B., S.R. Moulton, and G.T. Lester. 2003. Determining the quality of taxonomic data.
Journal of the North American Benthological Society 22:621-631.
University of Florida Herbarium. 2009. Preparation of Plant Samples
for Deposit as Herbarium Vouchers, http://www.flmnh.ufl.edu/herbarium/voucher.htm,
Accessed June 12, 2009.
USEPA. 2002. Methods for Evaluating Wetland ConditionL #10 Using Vegetation to Assess
Environmental Conditions in Wetlands. EPA-822-R-02-020, Office of Water, U.S.
Environmental Protection Agency, Washington, DC.
USEPA. 2006. Survey of the Nation's Lakes. Laboratory Methods Manual. EPA841-B-06-005.
U.S. Environmental Protection Agency, Washington, DC.
USEPA. 2006. Survey of the Nation's Lakes. Quality Assurance Project Plan. EPA841-B-07-
003. U.S. Environmental Protection Agency, Washington, DC.
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National Wetlands Condition Assessment
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5. SOIL
5.1 Introduction to Indicator
The presence of hydric soil is a defining characteristic of wetland ecosystems, and both
influences and is influenced by hydrology and vegetation (Tiner 1999, Mitsch and Gosselink
2007). Categories of data to be collected include the following:
1. Hydric soil field indicators
2. Description of soil profile and physical characteristics
3. Soil chemistry
4. Soil disturbance
For more detailed information please see the report Ecological Indicators for the 2011 National
Wetland Condition Assessment (in preparation).
5.1.1 References
Mitsch, W.J., and J.G. Gosselink. 2007. Wetlands. John Wiley & Sons, Hoboken, NJ.
Tiner, R. W. 1999. Wetland Indicators: A Guide to Wetland Identification, Delineation,
Classification, and Mapping. Lewis Publishers, CRC Press, Boca Raton, FL, USA.
5.2 Quality Assurance/Quality Control
Standardized lab protocols, consistent training of all lab technicians, lab assistance visits to all
labs, and availability of experienced technical personnel to respond to site-specific questions as
they arise are important to ensuring the quality of lab data. Additionally, control measures to
minimize measurement error among lab technicians and laboratories include the use of a
Control Sample, a Blank Sample, Data Review, and Data Validation (QAPP section 5.5.6).
A Control Sample represents a sample of known concentration for a particular attribute. A
Control Sample is collected in bulk for an attribute and repetitively analyzed to determine
statistical control limits (i.e., range of expected values) for the particular method. A Control
Sample is analyzed in conjunction with every batch of samples to ensure the method was run
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correctly. If the value of the Control Sample falls outside the expected range of values then the
process has failed and the batch is flagged for reanalysis.
A Blank Sample is used to ensure equipment is thoroughly cleaned before each use. A Blank
Sample is especially important when measuring soil chemistry (i.e., trace metals) because
concentrations may be quite small. A Blank Sample is analyzed in conjunction with every batch
of samples to ensure that proper equipment cleaning protocols are followed. If the value of the
Blank Sample does not equal zero or fall below the method detection limit, then the equipment
is not clean and the batch is flagged for reanalysis.
The process of Data Validation is described here. Laboratory data undergo four Data
Reviews, first by the Bench Analysts, second by the Lead Analyst, third by the Project
Coordinator Soil Scientist, and fourth by a Soil Scientist Liaison with expertise in soils from the
region where the samples are from. The Bench Analysts verifies that blank and control samples
return results that fall within established control limits. The Lead Analyst examines the data for
inconsistencies and apparent anomalies; inconsistencies usually take the form of unexpected
high or low values for a particular analyte or values that do not fit with the expected trend of a
soil profile. The Project Coordinator will use professional judgment to determine whether the
project data are self-consistent and congruent with the site data collected in the field;
incongruities within the data that can be explained either by site data or the results of other
analytes are recorded. A final review is given by a Soil Scientist Liaison to the area of sample
origin, before the data are released.
5.3 Receiving Regulated Soils
Soils that may contain pests (i.e., bacteria, plant viruses, fungi, nematodes, and life stages of
destructive mollusks, acari, and insects) are regulated by U.S Department of Agriculture's
Animal and Plant Health Inspection Service (APHIS). Areas within states that are under Federal
quarantine must follow the conditions and safeguards prescribed by APHIS before shipping to
another part of the country. To ensure that the NWCA is in compliance with APHIS
recommendations, all soils collected for the survey will be shipped as regulated soils.
Participating labs are responsible for obtaining and maintaining a valid permit for receiving
regulated soils (see example, USDA APHIS PPQ 525-A, Figure 5-1 below).
Upon arrival at the lab soil samples will be separated into regulated and non-regulated based on
their county and state of origin (as recorded on the water proof label affixed to the outside of the
sample bag). The lab is responsible for following all APHIS protocols when handling or
disposing regulated soils as found in 7 CFR 330.300.
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APHIS Animal and
-^^ Plant Health
^^F Inspection Service
United States Department of Agriculture
Animal and Plant Health Inspection Service
4700 River Road
Riverdale, MD 20737
Permit to Receive Soil
Regulated by 7 CFR 330
This permit was generated electronically via the ePermits system.
PERMITTEE NAME: Dr. Thomas Reinsch
COMPANY: USDA-NRCS-NSSC
RECEIVING .ADDRESS: Federal Building. Room 152, MS 41
100 Centennial Mall North
Lincoln. NE 68508-3866
Federal Building. Room 152, MS 41
100 Centennial Mall North
Lincoln. NE 6S50S-3866
(402)437-4179
(402) 437-5760
PERMIT NUMBER: P330-08-00009
APPLICATION NUMBER: P525-071002-007
DATE ISSUED: 01 14/2008
MAILING .ADDRESS:
PHONE:
FAX:
EXPIRES:
01/14/2011
PORTS OF ARRIVAL/PLANT INSPECTION STATIONS: Various Ports of Entry Staffed by CBP-Agriculture
Inspection
HAND CARRY: No
Under the condition? specified, this permit authorizes the following:
Quantity of Soil per Shipment and Treatment
Over 3 Ibs
PERMIT CONDITIONS
1. This permit authorizes the importation of soil, under the conditions specified below. Upon arrival in the United
States, the articles, shipping contamer(s). and paperwork are subject to inspection by officials of Customs and
Border Protection. Agriculture Inspection (C'BP-AI) and the USDA. Plant Protection and Quarantine (PPQ).
2. Under the Plant Protection Act. individuals or corporations who fail to comply with the following conditions and
authorizations, or who forge, counterfeit, or deface permits or shipping labels will receive civil or criminal penalties,
and will have all current permits cancelled and future permit applications denied.
3. Any person who unloads, lands, or otherwise brings or moves into or through the United States any regulated
plants, plant products, plant pests, soil or other products or articles in violation of the regulations will be subject to
prosecution under the applicable provisions of the law.
THIS PERMIT HAS BEEN APPROVED ELECTRONICALLY BY THE FOLLOWING
PPQ HEADQUARTER OFFICIAL VIA EPERMITS.
Maria Corpuz
DATE
01/14/2008
'.VARXJN'G: Any alte-adcm. former.' or unauthorized use of thi; Federal Form is sub'ect to civil penalties of up to $250.000 (/ U.S.C s .•' .'
510,000, or imprisonment of not more -Jiau 5 years, or both (IS U.S.C.s 1001)
i or piurishable bv :< fir.e of net raore -J
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APHB Animal and <^^30S^ PI
4. All foreign cargo of agricultural interest is inspected at the first port of arrival or the first port of unlading. If a
shipment arrives at a port without the required official personnel available to do the proper inspection, aud'br
treatment, any subsequent movement, or any transfer and or transloading. must be approved by CBP-AI
5. A copy of this permit must accompany all shipments authorized under this permit.
6. The soil is to be shipped in sturdy, leak-proof containers.
7. CBP-AI and PPQ have the option to order and approve treatment, re-exportation or destruction of a shipment, a
portion of a shipment, or any other material associated with the shipment (i.e. pallets, packaging, means of
conveyance). This will be done if the official personnel find that the shipment requires treatment as a condition of
entry, is contaminated with a quarantine plant pest or pests, is commingled with prohibited plant material, or if
required documentation is incomplete or missing.
8. The shipment must be free from foreign matter or debris, plants, noxious weed seeds, and living organisms such
as parasitic plants, pathogens, insects, snails, and mites. Material found to be commingled with unauthorized material
will be subject to the same action (i.e. re-export, destruction) as the unauthorized material.
9. All solid wood packing material (SWPM) present with this shipment must be in compliance with ISPM 15
treatment and IPPC stamp requirements and enforcement. Noncomphant shipments will be treated, re-exported or
destroyed at the consignee's expense.
10. All costs and arrangements for the safeguarding of the cargo and the transportation of the cargo are the
responsibility of the importer, broker, or other parties associated with the shipment.
11. The shipment can be leleased without treatment at the port of entry to the permittee's address listed on the permit
or label, or an authorized user only if the final destination is an approved facility listed at
https:/ webOl.aphis.usda.gov PPQ AuthSoilLabs.usf•1web?openfoim.
12. Permit is to be utilized by the permittee or authorized user only (authorized users must present a written, dated.
and signed statement on letterhead from the permittee, along with a valid ID and a copy of this permit).
13. There is no further distribution of soil without prior approval from the State and Federal Regulatory Officials
Soil is to be used strictly for analysis in a laboratory environment at USDA-NRCS-NSSC located in Lincoln. ME.
14. Upon receipt, all samples will remain within the approved soil laboratory identified on this permit. Laboratory
access is restricted to individuals authorized by the permit holder.
15. Tins permit does not authorize the use of soil for growing purposes and or the isolation or culture of organisms
sourced from imported soil.
16. All unconsurned soil, containers, and effluent is to be autoclaved. incinerated, or properly sterilized by the
permittee at the conclusion of the project as approved and prescribed by PPQ in the compliance agreement.
17. Valid for shipments of soil not heat treated at the port of entry, only if a Compliance Agreement (PPQ Form
519) has been completed and signed. Compliance Agreements and Soil Permits are non-transferable. Notify local
USDA office promptly if the permittee leaves the company.
IS. Tins permit authorizes shipments from all foreign sources, including Guam. Hawaii-
Puerto Rico, and the U.S. Virgin Islands through any U.S. port of entry.
END OF PERMIT CONDITIONS
Permit Number P330-CB-C0009
THIS PERMIT HAS BEEN .APPROVED ELECTRONICALLY BY THE FOLLOWING
PPQ HEADQUARTER OFFICIAL VTA EPERMTTS.
^~~ CT-ZT
Maria Corpuz
DATE
01/14/2008
WARNING: Any alteradon. forgery or unauthorized use of this Federal Form is subject to civil penalties of up to $250,000 (7 U.S.C.s 7734(b» or punishable by a tine of not more i
510,000. or itnpnsoiuiieut of tict more iban 5 years, or both (18 U.S.C.s 1001)
Figure 5-1: USDA APHIS PPQ 525-A for receiving regulated soils.
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5.4 Parameters for the NWCA
A total of 15 parameters will be measured from each soil sample collected (Table 5-1). Table 5-
1 also lists example methods that have been demonstrated to achieve the required objectives.
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Table 5-1: Soil parameters measured for the National Wetlands Condition Assessment.
Method
PSDA, <2mm, air
dry
Calcium carbonate
equivalent, <2mm
Calcium carbonate
equivalent,
<20mm
Total Carbon,
Nitrogen and
Sulfur
PH
Cation exchange
capacity
Ammonium
Oxalate Extraction
SSL Method
SAlala
4Elalalal
4Elalala2
4H2al-3
4Cla2ala-bl
4Cla2a2a-bl
4Blalbl-4
4G2alal-5
Analyte
Clay
Silt
Sand
CaCO3
CaCO3
C
N
S
1:1 H20
1'2 001 M
CaCI2
CEC
Ca2+
K+
Mg2+
Na+
Al
Fe
Mn
Units
%
%
%
%
%
%
%
%
PH
pH
cmol(+) kg"1
cmol(+) kg"1
cmol(+) kg"1
cmol(+) kg"1
cmol(+) kg"1
%
%
mg/kg
MDL
na
na
na
0.5
0.5
0.04
0.04
0.04
na
na
0.1
0.07
0.06
0.01
0.2
0.002
0.0001
0.1
PQL
na
na
na
2.5
2.5
0.2
0.2
0.2
na
na
0.6
0.4
0.3
0.07
1.0
0.009
0.0006
0.6
Reproducibility
Value
32
22
8.3
9.6
3.1
0.19
0.016
6.1
5.4
26
15
0
9
1
1.1
0.6
296
N
296
297
297
134
184
183
183
129
129
75
77
77
77
77
102
102
102
RL
2
3
2
1
0.3
0.04
0.02
0.3
0.3
2
1
0.1
0.6
0.3
0.2
0.08
42
Potential Sample
Range
0 to 93.1
0.1 to 100
0 to 94.5
nd* to 105
nd to 96
nd to 62.43
nd to 11. 193
nd to 21.86
2.4 to 10.5
2.3 to 10.5
nd to 252
nd to 507.3
nd to 17.4
nd to 147.1
nd to 650.3
nd to 15.62
nd to 20. 15
nd to 15730.7
Accuracy
Objective
0.01%
0.001%
0.01%
0.1 pH unit
0.1 pH unit
O.lcmol(+) kg-
1
O.lcmol(+) kg-
1
O.lcmol(+) kg-
1
O.lcmol(+) kg-
1
O.lcmol(+) kg-
1
0.01%
0.01%
1.0 mg/kg-1
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Laboratory Methods Manual
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Electrical
Conductivity
Dithionite-Citrate
Extraction
Olsen Phosphorus
Mehlich
Phosphorus
Trace Elements
4Flalalal
4Glal-3a-bl
4D5ala-bl
4D6ala-bl
4Hlalalal-
20
P
Si
EC
Al
Fe
Mn
P
P
Ag
As
Ba
Be
Cd
Co
Cr
Cu
Hg
Mn
Mo
Ni
P
Pb
Sb
Se
mg/kg
%
mmhos cm"1
%
%
%
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
Hg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
Hg/kg
26
0.0002
0.001
0.001
0.01
0.0006
0.1
0.1
0.001
0.002
"0"
0.001
0.001
"0"
0.006
0.002
1.3
0.002
0.001
0.009
0.4
0.001
0.002
1.8
129
0.001
0.005
0.006
0.07
0.003
0.7
0.6
0.01
0.01
"0"
0.01
0.01
"0"
0.03
0.01
6.50
0.01
0.01
0.05
2.07
0.01
0.01
9.00
867
0.41
3.1
0.19
2.2
0.091
13
19
1.8
50
500
0.86
1.6
10
22
78
212
1311
na
139
949
53
0.3
102
102
17
207
207
207
32
12
126
126
126
126
126
126
125
126
126
126
na
126
126
126
109
234
0.05
0.3
0.09
0.4
0.03
3
2
0.7
8
54
0.3
0.4
1
8
7
58
63
na
17
106
6
0.2
nd to 16926.4
ndto6.13
1.0 mg/kg-1
0.01%
0.01 mmhos
nd to 167.4 cm-1
ndto8.6 0.1%
ndto36.6 0.1%
ndto3.6 0.1%
nd to 399. 7 O.lmg/kg-1
nd to 1232.3 O.lmg/kg-1
nd to 175.62
nd to 1808.06
0.02 to 4415.8
nd to 29.98
nd to 85.68
nd to 1125.58
nd to 2020.31
nd to 1036.28
nd to 26060
nd to 692942
nd to 235. 17
tr* to 3347.36
nd to 70708.6
nd to 12287.4
nd to 42. 01
nd to 16523.1
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Bulk Density
3Bla
3Blb
3Blc
3Bld
Sn
Sr
V
W
Zn
Dbf
Db33
Dbod
Dbr
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
g/cm3
g/cm3
g/cm3
g/cm3
0.005
0.001
"0"
"0"
0.006
na
na
na
na
0.03
0.01
"0"
"0"
0.028
na
na
na
na
na
na
41
2.3
157
na
na
na
na
na
na
126
88
126
na
na
na
na
na
na
15
2
26
na
na
na
na
nd to 1117.66
nd to 10895
nd to 1064.65
nd to 137.39
0.06 to 10379.1
0.15 to 2.6
0.06 to 2.53
0.07 to 2.53
32 to 428.8
0.01 g/cm"3
0.01 g/cm"3
0.01 g/cm"3
0.01 g/cm"3
*nd= not detect
*tr= trace quantity
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5.5 Laboratory Sample Collection and Preparation
The purpose of any soil sample is to obtain information about a particular soil and its
characteristics. Sampling provides a means to estimate the parameters of these soil
characteristics with an acceptable accuracy at the lowest possible cost (Petersen and Calvin,
1986). Sub-sampling also may be used, as it permits the estimation of some characteristics of
the larger sampling unit without the necessity of measurement of the entire unit. Sub-sampling
reduces the cost of the investigation, but usually decreases the precision with which the soil
characteristics are estimated. Efficient use of sub-sampling depends on a balance between
cost and precision (Petersen and Calvin, 1986).
Soil variability and sample size are interferences to sample collection and preparation. The
objective of laboratory preparation is to homogenize the soil samples used in chemical,
physical, and mineralogical analyses. At each stage of sampling, an additional component of
variability, the variability within the larger units, is added to the sampling error (Petersen and
Calvin, 1986). Soil material needs to be adequate in amount and thoroughly mixed in order to
obtain a representative sample.
Bulk Samples: Laboratory identification numbers and preparation codes are assigned to bulk
soil samples. These identification numbers are unique EPA- and laboratory-assigned numbers
that carry important information about the soil sample (e.g., pedon, soil horizon, location, and
year sampled). Laboratory preparation codes depend on the properties of the sample and on
the requested analyses. These codes carry generalized information about the characteristics of
the analyzed fraction, i.e., the water content (e.g., air-dry, field-moist) and the original and final
particle-size fraction (e.g., sieved <2-mm fraction processed to 75 (im) and by inference, the
type of analyses performed. Identification numbers and preparation codes are reported on the
Primary Characterization Data Sheets.
All soils from quarantined areas are strictly controlled under APHIS quarantine regulations 7
CFR 330.
Natural Fabrics, Clods, and Cores: Laboratory identification number and preparation codes
are assigned to natural fabrics (NF), clods, and cores. These identification numbers typically
relate to a corresponding bulk sample.
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5.5.1 Summary of Method
For most standard chemical, physical, and mineralogical analysis, the field sample is air-dried,
crushed, and sieved to <2 mm. Field-moist, fine-earth fraction samples are processed by forcing
the material through a 2-mm screen by hand or with a large, rubber stopper and placed in a
refrigerator for future analysis. Depending on the nature of the soil material and requested
analyses, air-dry and/or field-moist samples may also be prepared as whole-soil samples or
processed further to finer fractions than <2 mm. Air-dry, <75 urn sub-samples for major and
trace elements are processed metal-free.
Generally, weight measurements are made and recorded on the 20- to 75-mm, 5- to 20-mm,
and 2- to 5-mm fractions, with these fractions then discarded. In some cases, these weight
measurements may not be recorded. Additionally, some or all of these >2-mm fractions may be
processed to a finer fraction and saved for chemical, physical, and mineralogical analysis. For
example, after the respective weights of the 5- to 20-mm and 2- to 5-mm fractions are recorded,
these fractions may be recombined and crushed to <2 mm in a laboratory jaw crusher, with the
recombined material saved for laboratory analysis. In other cases, the fine-earth fraction and
the >2-mm fractions are homogenized and passed through a laboratory jaw crusher to reduce
all material to pass a 2-mm sieve, with the processed material saved for laboratory analysis.
5.5.2 Interferences
Soil variability and sample size are interferences to sample collection and preparation. At each
stage of sampling, an additional component of variability, the variability among smaller elements
within the larger units, is added to the sampling error (Petersen and Calvin, 1986). Soil material
needs to be in adequate amount and thoroughly mixed to obtain a representative sample.
Soil is mixed by moving it from the corners to the middle of processing area and then by
redistributing the material. This process is repeated four times. Enough soil material needs to
be sieved and weighed to obtain statistically accurate rock fragment content. In order to
accurately measure rock fragments with a maximum particle diameter of 20 mm, the minimum
specimen size ("dry" weight) that needs to be sieved and weighed is 1.0 kg. Refer to ASTM
Standard Practice D 2488 (American Society for Testing and Materials, 2004). A homogenized
soil sample is more readily obtained from air-dry material than from field-moist material.
Whenever possible, "moist" samples or materials should have weights two to four times larger
than for "dry" specimens (American Society for Testing and Materials, 2004). Refer to ASTM
Standard Practice D 2488 (American Society for Testing and Materials, 2004).
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5.5.3 Safety
Dust from sample processing is a nuisance. A mask should be worn in order to avoid breathing
dust. Keep clothing and hands away from the crusher and pulverizer when these machines are
in use. Use a face-shield and goggles when operating the jaw crusher. Use goggles when
operating the air compressor. The HCI used to check carbonates can destroy clothing and
irritate skin. Immediately rinse acid with water from clothing or skin and seek professional
medical help, if needed.
5.5.4 Equipment
• Electronic Balance, ±1-g sensitivity and 15-kg capacity
• Cardboard trays for sample storage
• Trays, plastic, tared
• Sieves, square-hole, stainless steel
o 80 mesh, 180|im
o 10 mesh, 2 mm
o mesh, 4.75 mm
o 19 mm, 3/4 in
o 76 mm, 3 in
• 200-mesh, 75 urn
• Nylon cloth sieve
• 40-mesh, 0.425 mm
• Pulverizer
• Wooden rolling pin
• Rubber roller
• Laboratory jaw crusher, Retsch, Model BB2/A, Brinkmann Instruments Inc., Des Plaines,
IL
Metal plate, 76 x 76 x 0.5 cm
• Containers, paper, 12-oz, with lids
• Containers, plastic, 1-pint, 1, 4, and/or 8 oz with tops
• Scintillation glass vials, 20-mL
• Metal weighing cans, 2-oz
• Brown Kraft paper
• Air compressor, Cast-iron Series, SpeedAire, Campbell Hausfeld Mfg. Co., Harrison, OH.
• Planetary ball mill, Fritsch, Model P-5, Gilson, Lewis Center, OH
• Syalon balls, 12- to 15-mm, and bowls, 80-mL
• Metal weighing cans, 2-oz
• Cross beater mill, Retsch, Brinkmann Instruments Inc., Des Plaines, IL.
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5.5.5 Reagents
• Reverse osmosis (RO) water
1A/HCI
• Sodium hexametaphosphate solution. Dissolve 35.7 g of sodium hexametaphosphate
(NaPO3)6 and 7.94 g of sodium carbonate (Na2CO3) in 1 L of RO water.
5.5.6 Procedures
1. Weigh soil sample in sample bag to nearest g when logged-in and record weight.
2. Weigh sample in sample bag to nearest g before air-drying and record weight.
3. Remove soil sample from sample bag and distribute on a plastic tray. Thoroughly mix soil
material.
4. Before air-drying, weigh sample on a tared tray (tray weight) to nearest g and record weight.
5. Air-dry the sample in an oven at 30 to 35° C for 3 to 7 days.
6. Weigh sample to nearest g after air-drying and record weight.
7. Weigh sample to nearest g and record weight. This weight includes the >2-mm fractions.
8. Roll soil material on a flat, metal plate that is covered with brown Kraft paper with wooden
rolling pin and/or rubber roller to crush clods to pass a 2-mm sieve. For samples with easily
crushed coarse fragments, substitute a rubber roller for a wooden rolling pin. Roll and sieve
until only the coarse fragments that do not slake in sodium hexametaphosphate solution
remain on sieve. Crush clayey soils that contain no coarse fragments in a laboratory jaw
crusher.
9. Process air-dry soil by sieving to <2-mm. Thoroughly mix material by moving the soil from
the corners to the middle of the processing area and then by redistributing the material.
Repeat process four times. For preparation of the >2-mm fractions, see below.
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10. Procedures for air-dry, <2-mm fractions include but are not limited to the following:
a. Standard Analyses: For standard chemical, physical, and mineralogical
analysis, select material for representative sub-samples from at least five
different areas on the plastic tray. Prepare one sub-sample of the air-dry, sieved
<2-mm fraction in a 12-oz paper container.
b. Salt Analyses: For a saturation paste when salt analyses are requested, select
material for representative sub-samples from at least five different areas on the
plastic tray. Prepare one sub-sample of the air-dry, sieved <2-mm fraction in a 1-
pint plastic container.
c. 1500-kPA: For air-dry 1500-kPa water content of mineral and organic materials,
select material for representative sub-samples from at least five different areas
on the plastic tray. Prepare a sub-sample of air-dry, sieved <2-mm fraction in a
1-oz plastic cup.
d. Air-Dry/Oven-Dry Ratio: For air-dry/oven-dry (ADOD) ratio (required if any air-
dry analysis are determined), select material for representative sub-samples from
at least five different areas on the plastic tray. Prepare a sub-sample of the air-
dry, sieved <2-mm fraction in a 2-oz metal weighing can.
e. Presence of Carbonates: Use a sub-sample of the ADOD sample (procedure
1B1 b2b4) and check for the presence of carbonates. Reference samples
(knowns) are available for comparisons. Place 1 g of the air-dry fine-earth
fraction in porcelain spot plate, add reverse osmosis water, and stir to remove
entrapped air. Add 1 N HCI to soil, observe amount of effervescence, and record
as follows:
i. None - No visual efferevescence.
ii. Very Slight - Bubbles rise at a few points in the sample and consistently
appear at the same point in either a steady stream of tiny bubbles or in a
slower stream of larger bubbles. Do not mistake trapped air bubbles for a
positive test. Generally, these air bubbles appear immediately after the
addition of 1 N HCI.
iii. Slight - More small bubbles, and possibly a few larger bubbles, appear
throughout the sample than with a very slight reaction.
iv. Strong - More large bubbles are evident than with a slight reaction. Often
the reaction is violent at first and then quickly decreases to a reaction that
produces many small bubbles.
v. Violent - The sample effervesces violently. Many large bubbles appear to
burst from the spot plate.
f. Total Carbon, Nitrogen, and Sulfur Analysis: For total C, N, and S analyses,
select material for representative sub-samples from at least five different areas
on the plastic tray. Prepare one sub-sample of the air-dry, sieved <2-mm fraction
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processed in a cross beater mill to (« 80 mesh, 180 |jm) in a 20-mL scintillation
glass vial.
g. Calcium Carbonate and Gypsum: Use the prepared sub-sample (procedure
1B1b2d1) for the determination of the amount of carbonates and/or gypsum.
h. Chemical Analysis of Organic Materials: For chemical analysis of organic
materials, select material for representative sub-samples from at least five
different areas on the plastic tray. Prepare a sub-sample of the air-dry, sieved <2-
mm fraction processed in a cross beater to a fine grind (« 80 mesh, 180 urn) in a
12-oz paper container.
i. Total Major and/or Trace Elements: For total major and/or trace element
analyses, select material for representative sub-samples from at least five
different areas on the plastic tray. Prepare one sub-sample of the air-dry, metal
free, sieved <2-mm fraction processed in a planetary ball mill for 2 min and
sieved to <75 urn (200 mesh) in a 20-mL scintillation glass vial.
11. The following procedures are used for samples with >2 mm fractions. These fractions
include mineral coarse fragments as well wood fragments that are >20 mm in cross section
and cannot be crushed and shredded with the fingers. If the >2-mm fractions are to be
weighed, recorded, and discarded, with no further laboratory analysis, proceed to procedure
a. When the data for >2-mm fractions are not recorded, proceed to procedure b. If the >2-
mm fractions contain carbonate- or gypsum-indurated material, and laboratory analysis is
requested, proceed to procedure c. If the >2-mm fractions are Cr or R material (Soil Survey
Staff, 1999), and laboratory analysis is requested, proceed to procedure d.
a. Particle-Size Analysis, Recorded: Process the air-dry soil by sieving to <2 mm
as described in step 9. In this procedure, weight measurements are made on the
20- to 75-mm, 5- to 20-mm, and 2- to 5-mm fractions. Weigh soil material with
diameters of 2 to 5 mm. If difficult to separate the <2-mm fraction from fragments,
soak (100 g of 2- to 5-mm fraction) in sodium hexametaphosphate solution for 12
h. Air-dry, weigh the material that does not slake, record weight, and discard.
Weigh, record weight, and discard particles with diameters of 20 to 75 mm and 5
to 20 mm. The <2-mm fraction is saved for chemical, physical, and mineralogical
analysis.
b. Particle-Size Analysis, Not Recorded: This procedure is the same as
described in step 9 except the weight of the >2-mm fractions are not recorded
and all analytical results are reported on a <2-mm basis.
c. Particle-Size Analysis, Recorded, 2- to 20-mm Fraction Processed to <2-
mm, Chemical, Physical, and Mineralogical Analysis: This procedure is
commonly used for samples with carbonate- or gypsum-indurated material.
Process the air-dry soil by sieving to <2 mm. Weigh soil material with diameters
of 20- to 75-mm, 5 to 20-mm, and 2 to 5-mm and record weights as described in
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procedure 11 a. The 5- to 20-mm and 2- to 5-mm fractions are then recombined
after their respective weights are recorded. The recombined, 2- to 20-mm,
material is crushed to <2 mm in a laboratory jaw crusher. This material is saved
for laboratory analysis and analytical results reported on the 2- to 20-mm basis.
The <2-mm material is also saved for chemical, physical, and mineralogical
analysis. If carbonate or gypsum accumulations are soft and easily pass a 2-mm
sieve, the standard procedure is usually requested.
d. Particle-Size Analysis, Not Recorded, Whole-Soil Processed to <2-mm,
Chemical, Physical, and Mineralogical Analysis: This procedure is mainly
used to prepare samples from Cr or R soil horizons (Soil Survey Staff, 1999).
Homogenize particles with diameters >2 mm and the fine-earth material (<2-mm)
and crushed to <2-mm in a laboratory jaw crusher. This material is saved for
laboratory analysis and analytical results reported on the whole-soil basis.
5.5.7 Calculations
Calculations for coarse fragments are reported in 3A2.
5.5.8 Report
Reported data include but are not limited to the following:
1. Weight (g) of field-moist soil sample
2. Weight (g) of air-dry soil sample
3. Weights (g) of processed air-dry soil
4. Weight (g) of 20- to 75-mm fraction
5. Weight (g) of 5- to 20-mm fraction
6. Weight (g) of 2- to 5-mm fraction
7. Weight (g) of sub-sample of 2- to 5-mm fraction before slaking
8. Weight (g) of sub-sample of 2- to 5-mm fraction after slaking
9. Effervescence with HCI (None, Very Slight, Slight, Strong, Violent)
5.5.9 Precision and Accuracy
Precision and accuracy data are available from the SSL upon request.
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5.5.10 References
American Society for Testing and Materials. 2004. Standard practice for description and
identification of soils (visual-manual procedure). D 2488.. Annual book of ASTM
standards. Construction. Section 4. Soil and rock; dimension stone; geosynthesis. Vol.
04.08. ASTM, Philadelphia, PA.
Bates, T.E. 1993. Soil handling and preparation, p. 19-24. In M.R. Carter (ed.) Soil sampling
and methods of analysis. Can. Soc. Soil Sci. Lewis Publ., CRC Press, Boca Raton, FL
Petersen, R.G., and LD. Calvin. 1986. Sampling, p. 33-51. In A. Klute (ed.) Methods of soil
analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr 9. ASA
andSSSA, Madison, Wl.
Soil Survey Staff. 1995. Soil survey laboratory information manual. Version No. 1.0. USDA-
NRCS. Soil Survey Investigations Report No. 42. U.S. Govt. Print. Office, Washington,
DC.
Soil Survey Staff. 1996. Soil survey laboratory methods manual. Version No. 3.0. USDA-NRCS.
Soil Survey Investigations Report No. 42. U.S. Govt. Print. Office, Washington, DC.
Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys. 2nd ed. Agric. Handb. No. 436. USDA-NRCS. U.S. Govt. Print.
Office, Washington, DC.
5.6 Sampling Archiving
All remaining soil from a sample will be archived for potential re-analysis in the future. The
remaining samples are air-dried and passed through a 2mm sieve (following procedures for
sample preparation in Section 5.5.6) before storage in a climate controlled facility. Air-drying will
stop most biological activity, but long-term storage may cause minor changes in concentrations
of organic C, N, and S. Therefore, re-analysis of these fractions would not be appropriate.
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5.7 PSDA Air-dry, <2mm Particles
5.7.1 Summary of Method
The standard SSL procedure for analysis of particles with <2-mm diameters is the air-dry
method (procedure 3A1a1a), whereby a 10-g sample is pretreated to remove organic matter
and soluble salts. The sample is dried in the oven to obtain the initial weight, dispersed with a
sodium hexametaphosphate solution, and mechanically shaken. The sand fraction is removed
from the suspension by wet sieving and then fractionated by dry sieving. The clay and fine silt
fractions are determined using the suspension remaining from the wet sieving process. This
suspension is diluted to 1 L in a sedimentation cylinder, stirred, and 25-mL aliquots removed
with a pipet at calculated, predetermined intervals based on Stokes' law (Kilmer and Alexander,
1949). The aliquots are dried at 110°C and weighed. Coarse silt is the difference between
100% and the sum of the sand, clay, and fine silt percentages.
5.7.2 Interferences
The sedimentation equation that is used to measure the settling rates of particles of different
sizes is as follows:
where:
v = Velocity of fall
r = Particle radius
g = Acceleration due to gravity
ps = Particle density
pi = Liquid density
H = Fluid viscosity
This formula results from an application of Stokes' law and is referred to as Stokes' law.
Assumptions used in applying Stokes' law to soil sedimentation measurements are as follows:
1 . Terminal velocity is attained as soon as settling begins.
2. Settling and resistance are entirely due to the viscosity of the fluid.
3. Particles are smooth and spherical.
4. There is no interaction between individual particles in the solution (Gee and Bauder, 1986;
Gee and Or, 2002).
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Since soil particles are not smooth and spherical, the radius of the particle is considered an
equivalent rather than an actual radius. In this method, particle density is assumed to be 2.65 g
cc"1. Gypsum interferes with PSDA by causing flocculation of particles. Gypsum is removed by
stirring and washing the soil with reverse osmosis water. This procedure is effective if the soil
contains <25% gypsum. Partial flocculation may occur in some soils if excess H2O2 is not
removed from the soil after its use in organic matter oxidation. Treatment of micaceous soils
with H2O2 causes exfoliation of the mica plates and a matting of particles when dried in the
oven. Since exfoliation occurs in these soils, a true measurement of fractions is uncertain
(Drosdoff and Miles, 1938).
5.7.3 Precautions
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses) when handling acid and H2O2. Mix acids in ventilated fume
hoods. Use sodium bicarbonate and water to neutralize and dilute spilled acids. Heat samples
for removal of organic matter or cementing agents in ventilation hoods. Handle heated samples
with leather gloves. Perform the transfer of acid to gelatin capsules near a sink in case of
leakage or spills. Users should be familiar with centrifuge operation. Opposite centrifuge
bottles need to be balanced. Centrifuge should not be opened until centrifuge rotor has
completely stopped.
5.7.4 Equipment/Materials
• Beakers, 300 ml_, tared to 1 mg
• Ceramic filter candles, 0.3 urn absolute retention
• Rack to hold ceramic filter candle and sample container.
• Mechanical shaker, horizontal, 120 oscillations min-1, 1 1A in strokes, Eberbach 6000,
Eberbach Corp., Ann Arbor, Ml
• Cylinders, 1 L, white line fused onto glass at 1-L mark
Oven, 110°C
Hotplate, 100°C
Vacuum, 0.8 bars (SOkPa)
• Thermometer, 0 to 150°C
• Desiccator
• Motor driven stirrer, (Kilmer and Mullins, 1954)
• Hand stirrer, perforated disk fastened to a rod
• Adjustable pipet rack (Shaw, 1932; Fig. 1-3)
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Lowy pipets, 25 ml_, with overflow bulb
Polyurethane foam, pipe insulation that fits snugly around cylinder.
Sieve shaker with 12.7-mm (1/2 in) vertical and lateral movement at 500 oscillations min-1.
Accommodates a nest of 76-mm (3-in) sieves.
Weighing bottles, 90 ml_, with screw caps, tared to 1 mg
Weighing bottles, 90 ml_, tared to 1 mg
Weighing bottles, 90 ml_, tared to 0.1 mg
Drying dishes, aluminum
Timer or clock with second hand
Electronic balance, ±0.10-mg sensitivity
Electronic balance, ±1.0-mg sensitivity
Watch glass, 50- and 65-mm diameters
Evaporating dish, porcelain, 160-mm diameter, 31-mm height, with lip
Set of 76-mm (3 in) sieves, square weave phosphor bronze wire cloth except 300 mesh
which is twilled weave. U.S. series and Tyler Screen Scale equivalent designations are as
follows:
Sand Size
VCS
CS
MS
FS
VFS
Opening
(mm)
1.0
0.5
0.25
0.105
0.047
U.S. No.
18
35
60
140
300
Tyler Mesh
Size
16
32
60
150
300
Centrifuge, International No. 11, with No. 949 rotor head, International Equip. Co., Boston,
MA
Centrifuge bottle, 500 ml_
Torsion balance
Manometer, hand-held gauge and differential pressure, PCL-200A/C Series, Omega
Engineering, Stamford, CT.
Gelatin capsules, 5 ml_
Threaded weighing bottles, 90 mL
Machined PVC caps for threaded 90-mL weighing bottles, 3.2-cm (1 1/4 in) diameter with
1.1-cm (7/16 in) diameter hole drilled in center, O-ring seal
O-rings, 3.2 x 38.1 mm (1/8 x 1 1/2 in)
Septa, rubber, 7.9-mm (5/16 in) diameter. Place in machined cap.
Hypodermic needle, 25.4 mm (1 in), 23 gauge
Ultrasonic probe, 19-mm (3/4 in) horn, 20 kHz, 300 watts
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5.7.5 Reagents
• Reverse osmosis (RO) water, ASTM Type III grade of reagent water
Hydrogen peroxide (H2O2), 30 to 35%
• Sodium hexametaphosphate ((NaPO3)6), reagent grade
• Sodium carbonate (Na2CO3), reagent grade
• Sodium hexametaphosphate solution. Dissolve 35.7 g of (NaPO3)6 and 7.94 g of Na2CO3
in 1 L of RO water. See Section 7.12 for standardization of sodium hexametaphosphate
solution.
• Ethyl alcohol
• Calcium sulfate (anhydrous) or equivalent desiccant
• Hydrochloric acid (HCI), 6 N, technical grade. Dilute 1 L of concentrated HCI with 1 L of
RO water
• Sodium carbonate (Na2CO3) reagent. Dissolve 10.6 g Na2CO3 in RO water and make to 1
L(10mgCaCO3)
• 1 N sodium acetate (NaOAc) solution, buffered to pH 5. Dissolve 680 g of NaOAc in 4 L of
RO water. Add ~ 250 ml_ of acetic acid. Make to 5-L volume with RO water.
Sodium citrate solution, 0.3 M Na3C6H5O7'2H2O (88.4 g L-1)
• Sodium bicarbonate buffer solution, 1 M NaHCO3 (84 g L-1)
• Sodium dithionite (Na2S2O4 - hydrosulphite)
Saturated NaCI solution (solubility at 20°C; 360 g L-1)
• Sodium hydroxide solution (NaOH), 0.1 N. Dissolve 4 g NaOH pellets in 1 L of RO water.
5.7.6 Procedure
1. Weigh 10g of <2-mm, air-dry soil to nearest mg on an electronic balance and place into a
numbered, tared 300-mL, beaker. Wash and tare these beakers once every two months. A
quality control sample is included in each batch (<24 samples).
2. Add « 50mL of RO water and 5 to 7.5 mL of H2O2 to the soil sample at ambient temperature.
Cover the soil sample with a 50-mm watch glass. Allow initial oxidation of organic matter to
complete and then place sample on hot plate. If the froth from the reaction exceeds the
capacity of the beaker, transfer the sample to a larger beaker.
3. Place the sample on a hot plate and heat to 90°C. Add four 5 to 7.5-mL increments of H2O2
at 30-min intervals. Add additional H2O2 until organic matter oxidation is complete. Heat the
sample for an additional 45 min to decompose excess H2O2. If the reaction is violent, do
one or any combination of the following: (a) add small increments of ethyl alcohol to the
sample; (b) remove the sample from the hot plate to slow the reaction; (c) transfer sample to
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a 1000-mL beaker; or (d) reduce the amount of H2O2 to sample. Record any unusual
sample reactions.
4. Place the sample on the filter rack. Add 150 mL of RO water. Insert a filter candle, connect
to the vacuum trap assembly with tubing, and turn on the vacuum. Aspirate until liquid is
removed and only slightly dampened sample remains. Wash the sample four additional
times with « 150ml_ of RO water. Stir the sample with filter candle to ensure all soil particles
will be rinsed. During aspiration, it may be necessary to occasionally apply back-pressure to
filter candle and remove build-up of soil which inhibits aspiration. If the sample contains
gypsum and flocculates, then the following additional washings may be used. If the sample
contains 1 to 5% gypsum, stir the sample with a magnetic stirrer for 5 min and wash 5 times
with ~ 250ml_ of RO water each time. If the sample contains >5% gypsum, place the
sample in a 1000-mL beaker and stir the sample with a magnetic stirrer for 5 min then wash
5 times with « 750 mL of RO water each time to remove soluble gypsum.
5. Place sample in oven. Dry the sample overnight at 110°C. Remove the sample from the
oven, place in a desiccator, and cool to ambient temperature.
6. Record the total weight (TW) of the sample to the nearest mg.
7. Add the exact volume of sodium hexametaphosphate solution (« 10mL), equivalent to
0.4408g of sodium hexametaphosphate, to each sample. Subtract the weight of the sodium
hexametaphosphate (DW) that is contained in the extracted aliquot from the silt and clay
weights to calculate silt and clay percentages. To determine the exact volume of sodium
hexametaphosphate to add to each sample, refer to Section 5.7.6, step 9. Let stand until
sample is completely moistened by sodium hexametaphosphate. Add « 175mL of RO
water.
8. A sodium hexametaphosphate standardization is performed with each new batch of solution.
Use only designated weighing bottles for standardization. Wash and tare these bottles after
each standardization. Add duplicate aliquots (8.5, 9.0, 9.3, 9.6, 10.0, 10.3, 10.6, 11.0 mL) of
sodium hexametaphosphate solution to numbered tared, 90-mL weighing bottles. Oven-dry
aliquots overnight and record dry residue weight of sodium hexametaphosphate.
9. Determine the exact volume of sodium hexametaphosphate to add to each sample by
regressing the volume of sodium hexametaphosphate against the dry residue weight of
sodium hexametaphosphate and then by predicting the volume needed to dispense 0.4408g
of sodium hexametaphosphate into each sample.
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10. Place the sample in a horizontal shaker set at 120 oscillations min-1 and shake for 15 hours
(overnight).
11. Remove the sample from the shaker and pour through a 300-mesh (0.047-mm) sieve
mounted on a ring stand. Finger-rubbing of sample may be required during transfer to
speed washing of sample. Place a funnel below the sieve and a 1-L cylinder below the
funnel. Collect the silt and clay in the 1-L cylinder. Avoid using jets of water in washing the
sample. Wash and rub all particles from the beaker into the sieve. Continue to wash until
the suspension volume in the cylinder is ~ SOOmL. Sand and some of the coarse silt remain
on the sieve. Rinse all <20-um particles into the cylinder. Fill the cylinder to 1L and cover
with a 65-mm watch glass. Place pipe insulation around sample and blank cylinders to
prevent rapid changes in temperatures. Prepare a RO water blank to measure temperature
fluctuations. Allow the cylinder to stand overnight to equilibrate the suspension with the
room temperature. Wash the sand into an evaporation dish and dry the sand at 110°C
overnight.
12. Transfer the dried sand to a nest of sieves that has a top-to-bottom order of 1.0, 0.5, 0.25,
0.1, and 0.047 mm. Shake the sand for 3 min on a shaker that has 1.3-cm vertical and
lateral movements and oscillates at 500 strokes min-1. Record the weight of each separate
sand fraction (SWi) to the nearest mg. If optical analysis is requested, place the very fine
sand and fine sand fractions in gelatin capsules and the remaining sand fractions in a
labeled vial. Store capsules in the labeled vial. Wash sand dishes after every use.
13. Determine the percentage of fine silt and clay gravimetrically by removing an aliquot from
the suspension in the 1-L cylinder with a Lowy, 25-mL pipet. Periodically, gravimetrically
calibrate the delivery volume of the pipet by weighing the amount of RO water dispensed
from the pipet. Record the delivery volume (DV) and use the value to calculate the results.
Regulate the vacuum such that the pipet fills in «12 s. Record temperature (T1) of blank.
Mount the pipet on an adjustable pipet rack (Shaw, 1932). Stir the silt and clay suspension
with mechanical stirrerfor at least 5 min. Place the cylinder on a stable, vibrationless table
and stir with a hand stirrer in an up-and-down motion for 30s. Timing is started upon
completion of the stirring. Record the time that stirring is stopped. For the <20-um fraction,
slowly lower the closed pipet to a 10-cm depth in the suspension, turn on the vacuum, and
withdraw an aliquot at the calculated time (Table 1). Dispense the aliquot into a tared and
numbered, 90-mL weighing bottle. Rinse the pipet twice with RO water and dispense into
the tared, weighing bottle with the aliquot. For the <2-um fraction, pipet after a time of 4.5,
5, 5.5, or 6.5 h. Record temperature (T2) of blank. Use the average of T1 and T2 and
adjust the pipet depth in the suspension as indicated in Table 2. Repeat the procedure
described for the <20-um fraction. If determination of carbonate is required, use weighing
bottle with screw threads. Dry the aliquots at 110°C overnight and cool in a desiccator that
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contains calcium sulfate or an equivalent desiccant. Record the weight of the residue (RW)
to the nearest 0.1 mg.
14. Use the 90-mL, round-bottomed, weighing bottles for the <20-um aliquots. Wash and tare
after every fourth use. Use the 90-mL, square-bottomed, weighing bottles for the <2-um
aliquots. Wash and tare after every use.
5.7.7 Reporting
Report each particle-size fraction to the nearest 0.1 percent.
5.7.8 References
Bouyoucos, G.B. 1929. The ultimate natural structure of soil. Soil Sci. 28:27-37.
Drosdoff, M., and E.F. Miles. 1938. Action of hydrogen peroxide on weathered mica. Soil Sci.
46:391-395.
El Swaify, S.A. 1980. Physical and mechanical properties of Oxisols. p. 303-324. In B.K.G.
Theng (ed.) Soils with variable charge. N.Z. Soc. Soil Sci., Lower Hutt, N.Z.
Espinoza, W., R.H. Rust, and R.S. Adams Jr. Characterization of mineral forms in Andepts from
Chile. Soil Sci. Soc. Am. Proc. 39:556-561.
Gee, G.W., and J.W. Bauder. 1986. Particle-size analysis, p. 383-411. In A. Klute (ed.) Method
of soil analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9.
ASA and SSSA, Madison, Wl.
Gee, G.W., and D. Or. 2002. Particle-size analysis, p. 255-293. In J.H. Dane and G.C. Topp
(eds.) Methods of soil analysis. Part 4. Physical methods. Soil Sci. Am. Book Series No.
5. ASA and SSSA, Madison, Wl.
Grossman, R.B., and J.L. Millet. 1961. Carbonate removal from soils by a modification of the
acetate buffer method. Soil Sci. Soc. Am. Proc. 25:325-326.
Jackson, M.L. 1969. Soil chemical analysis—advanced course. 2nd ed. Univ. Wisconsin,
Madison.
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Kilmer, V.J., and L.T. Alexander. 1949. Methods of making mechanical analyses of soils. Soil
Sci. 68:15-24.
Kilmer, V.J., and J.K. Mullins. 1954. Improved stirring and pipeting apparatus for mechanical
analysis of soils. Soil Sci. 77:437-441.
Kubota, T. 1972. Aggregate formation of allophonic soils; Effects of drying on the dispersion of
soils. Soil Sci. and Plant Nutr. 18:79-87.
Maeda, T., H. Takenaka, and B.P. Warkentin. 1977. p. 229-263. Physical properties of
allophone soils. In N.C. Brady (ed.) Adv. Agron. Acad. Press, Inc., New York, NY.
Mehra, O.P., and M.L. Jackson. 1960. Iron oxide removal from soils and clays by a dithionite-
citrate system buffered with sodium bicarbonate, p. 237-317. In Clays and clay minerals.
Proc. 7th Conf. Natl. Acad. Sci. Natl. Res. Counc. Pub., Washington, DC.
Mikhail, E.H., and G.P. Briner. 1978. Routine particle size analysis of soils using sodium
hypochlorite and ultrasonic dispersion. Aust. J. Soil Res. 16:241-244.
Rabenhorst, M.C., and L.P. Wilding. 1984. Rapid method to obtain carbonate-free residues from
limestone and petrocalcic materials. Soil Sci. Soc. Am. J. 48:216-219.
Saly, R. 1967. Use of ultrasonic vibration for dispersing of soil samples. Sov. Soil Sci. 11:1547-
1559.
Shaw, T.M. 1932. New aliquot and filter devices for analytical laboratories. Ind. Eng. Chem.
Anal. Ed. 4:409.
Shields, L.G., and M.W. Meyer. 1964. Carbonate clay: Measurement and relationship to clay
distribution and cation exchange capacity. Soil Sci. Soc. Am. Proc. 28:416-419.
Skopp, J. 1992. Concepts of soil physics. University of Nebraska, Lincoln, NE.
Soil Survey Division Staff. 1993. Soil survey manual. USDA. Handb. No. 18. U.S. Govt. Print.
Office, Washington, DC.
Soil Survey Staff. 1996. Soil survey laboratory methods manual. Version No. 3.0. USDA-NRCS.
Soil Survey Investigations Report No. 42. U.S. Govt. Print. Office, Washington, DC.
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Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys. 2nd ed. Agric. Handb. No. 436. USDA-NRCS. U.S. Govt. Print.
Office, Washington, DC.
Sumner, M.E., 1992. The electrical double layer and clay dispersion, p. 1-32. In M.E. Sumner
and B.A. Stewart (eds.) Soil crusting, chemical, and physical processes. CRC Press Inc.,
Boca Raton, FL
Watson, J.R. 1971. Ultrasonic vibration as a method of soil dispersion. Soil Fertil. 34:127-134.
Williams, D.E. 1948. A rapid manometric method for the determination of carbonate in soils. Soil
Sci. Soc. Am. Proc. 13:127-129.
5.8 PS DA Pi pet Analysis, Particles <2 mm
5.8.1 Summary of Method
Standard SSL PSDA: The standard SSL procedure for analysis of particles with <2-mm
diameters is the air-dry method (procedure 3A1a1a), whereby a 10-g sample is pretreated to
remove organic matter and soluble salts. The sample is dried in the oven to obtain the initial
weight, dispersed with a sodium hexametaphosphate solution, and mechanically shaken. The
sand fraction is removed from the suspension by wet sieving and then fractionated by dry
sieving. The clay and fine silt fractions are determined using the suspension remaining from the
wet sieving process. This suspension is diluted to 1L in a sedimentation cylinder, stirred, and
25-mL aliquots removed with a pipet at calculated, predetermined intervals based on Stokes'
law (Kilmer and Alexander, 1949). The aliquots are dried at 110°C and weighed. Coarse silt is
the difference between 100% and the sum of the sand, clay, and fine silt percentages.
Fine-Clay Determination: The soil suspension from procedure 3A1a1a is used to determine
the fine-clay fraction. This suspension is stirred, poured into a centrifuge bottle, and centrifuged
at 1500 rpm. A 25-mL aliquot is withdrawn with a pipet. The aliquot is dried in the oven,
weighed, and the percentage of fine clay is calculated based on the total sample weight. The
time of centrifugation is determined from the following equation modified from Stokes' law
(Jackson, 1969).
tm = (63.0x108ri log (rs'1)) (Nm2 D^i2 Ap)'1
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where:
tm = Time in minutes
r| = Viscosity in poises
r = Radius in cm from center of rotation to sampling depth (3 cm + s)
s = Radius in cm from center of rotation to surface of suspension
Nm =rpm(1500)
Dp = Particle diameter in microns (0.2 urn)
Ap = Difference in specific gravity between solvated particles and suspension
liquid
63.0x108= Combination of conversion factors for convenient units of time in
minutes, tmin, Nm as rpm, and particle diameter in microns, Dp,.
Carbonate-Clay Determination: The residue from procedure is used to determine the
carbonate-clay fraction. This residue is treated with acid in a closed system. The pressure of
the evolved gas is measured. The pressure is related linearly to the CO2 content in the
carbonates. A manometer is used to measure the pressure.
Pretreatment and Dispersion Techniques: In the standard PSDA, an air-dry soil sample is
pretreated to remove organic matter and soluble salts. There are additional non-routine
chemical pretreatments for the removal of cementing agents that often prevent complete
dispersion. These pretreatments may be requested by the project coordinator as follows:
• Carbonate Removal: Carbonates are destroyed with a 1 N NaOAc solution buffered
to pH 5. The NaOAc solution is added to sample until carbonate bubbles no longer
evolve. The NaOAc solution is then washed from the sample. After destruction of
carbonates, the remainder of procedure is followed.
• Iron Removal: Soil samples are pretreated with H2O2 to remove organic matter.
Iron oxides are removed with bicarbonate-buffered, sodium dithionite-citrate solution
and heated until the sample color changes to a grayish color. The suspension is
flocculated with saturated NaCI solution and filtered to remove soluble salts. After
removal of iron oxides, the remainder of procedure is followed.
• Silica Removal: Soils are pretreated with H2O2 to remove organic matter. Soils with
Si cementation or coatings are pretreated with a weak NaOH solution overnight.
After removal of siliceous cementing agents, the remainder of procedure is followed.
• Ultrasonic Dispersion: A soil sample is pretreated to remove organic matter and
soluble salts. The sample is dried in the oven and weighed to obtain the initial
weight. Sodium hexametaphosphate solution is added to the sample and then made
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to 100-mL volume with RO water. The sample is subjected to ultrasonic vibration for
5 min. After dispersion with the ultrasonic probe, procedure is followed.
• Water Dispersible PSDA: Water dispersible particle-size distribution analysis may
also be determined from a soil suspension without the removal of organic matter or
soluble salts, or without the use of a chemical dispersant. Upon omitting these
procedural steps, the remainder of procedure is followed.
5.8.2 Interferences
Standard SSL PSDA: The sedimentation equation that is used to measure the settling rates of
particles of different sizes is as follows:
v = 2r2g(Ps-p,)/(9T1)
where:
v = Velocity of fall
r = Particle radius
g = Acceleration due to gravity
ps = Particle density
Pi = Liquid density
r| = Fluid viscosity
This formula results from an application of Stokes' law and is referred to as Stokes' law.
Assumptions used in applying Stokes' law to soil sedimentation measurements are as follows:
• Terminal velocity is attained as soon as settling begins.
• Settling and resistance are entirely due to the viscosity of the fluid.
• Particles are smooth and spherical.
• There is no interaction between individual particles in the solution (Gee and Bauder,
1986; Gee and Or, 2002).
• Since soil particles are not smooth and spherical, the radius of the particle is considered
an equivalent rather than an actual radius. In this method, particle density is assumed to
be 2.65 g cc"1.
• Gypsum interferes with PSDA by causing flocculation of particles. Gypsum is removed
by stirring and washing the soil with reverse osmosis water. This procedure is effective
if the soil contains <25% gypsum.
• Partial flocculation may occur in some soils if excess H2O2 is not removed from the soil
after its use in organic matter oxidation.
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• Treatment of micaceous soils with H2O2 causes exfoliation of the mica plates and a
matting of particles when dried in the oven. Since exfoliation occurs in these soils, a true
measurement of fractions is uncertain (Drosdoff and Miles, 1938).
Fine-Clay Determination: In the fine-clay determination, the distance from the center of
rotation to the surface of the suspension must be constant for each centrifuge bottle. The
particle density (pp) of the fine clay is assumed to be 2.5 g cc"1 (Jackson, 1969). The
suspension temperature must be used to enter the correct liquid viscosity in the equation.
Position the bottle under pipet without sudden movement of the centrifuge rotor, which causes
disturbance of solution. The withdrawal rate with pipet should be constant.
Carbonate-Clay Determination: The carbonate-clay analysis is semi-quantitative. It is
assumed that all of the carbonates are converted to CO2. This method measures all forms of
carbonates. In addition to Ca, the carbonates of Mg, Na, and K also react with the acid.
Analytical interferences may be caused by temperature changes within the reaction vessel. The
analyst should not touch the glass of the vessel when reading the pressure. When sealing the
vessel, the analyst should not hold onto the vessel any longer than necessary to tighten the cap.
The internal pressure must be equalized with the atmosphere. Approximately 3 to 5 s are
required to equalize the internal pressure of the bottle when piercing the septa with a needle.
The analyst should replace septa and O rings at regular intervals, as they develop leaks after
extensive use.
Pretreatment and Dispersion Techniques: The PSDA results are dependent on the
pretreatments used to disperse the soil. The presence of cementing agents such as
carbonates, Fe, and Si often prevent complete dispersion. In these cases, special pretreatment
and dispersion procedures may be performed upon request on either an air-dry or field-moist
sample. However, these special techniques in themselves may interfere with PSDA as follows:
• Carbonate Removal: The removal of carbonates with 1 N NaOAc (pH 5) results in
sample acidification. This pretreatment can destroy the primary mineral structure of clay
(Gee and Bauder, 1986).
• Iron Removal: If the temperature of the water bath exceeds 80° C during Fe removal,
elemental S can precipitate (Mehra and Jackson, 1960). This pretreatment can destroy
primary mineral grains in the clay fraction (El-Swaify, 1980).
• Silica Removal: The effects of Si removal with 0.1 N NaOH on the clay fraction and
particle-size distribution are unknown.
• Ultrasonic Dispersions): Ultrasonic dispersion has been reported to destroy primary
soil particles. Watson (1971) summarized studies that reported the destruction of biotite
and breakdown of microaggregates by ultrasonic dispersion. However, Saly (1967)
reported that ultrasonic vibration did not cause the destruction of the clay crystalline
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lattice or the breakdown of primary grains. The samples ranged from sandy to clayey
soils. The cementing agents represented humus, carbonates, and hydroxides of Fe and
Al. No standard procedures have been adopted using ultrasonic dispersion.
5.8.3 Safety
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses) when handling acid and H2O2. Mix acids in ventilated fume
hoods. Use sodium bicarbonate and water to neutralize and dilute spilled acids. Heat samples
in ventilated fume hoods for removal of organic matter or cementing agents in ventilation hoods.
Handle heated samples with leather gloves. Perform the transfer of acid to gelatin capsules
near a sink in case of leakage or spills.
Users should be familiar with centrifuge operation. Opposite centrifuge bottles need to be
balanced. Centrifuge should not be opened until centrifuge rotor has completely stopped
5.8.4 Equipment
Fleakers, 300 ml_, tared to 1 mg
Ceramic filter candles, 0.3 (im absolute retention (source currently unavailable)
Rack to hold ceramic filter candle and sample container.
Mechanical shaker, horizontal, 120 oscillations min-1, 1 1A in strokes, Eberbach 6000,
Eberbach Corp., Ann Arbor, Ml
Cylinders, 1 L, white line fused onto glass at 1-L mark
Oven, 110°C
Hotplate, 100°C
Vacuum, 0.8 bars (SOkPa)
Thermometer, 0 to 150°C
Desiccator
Motor driven stirrer, (Kilmer and Mullins, 1954)
Hand stirrer, perforated disk fastened to a rod
Adjustable pipet rack (Shaw, 1932; Fig. 1-3)
Lowy pipets, 25 ml_, with overflow bulb
Polyurethane foam, pipe insulation that fits snugly around cylinder.
Sieve shaker with 12.7-mm (1/2 in) vertical and lateral movement at 500 oscillations min-1.
Accommodates a nest of 76-mm (3 in) sieves.
Weighing bottles, 90 ml_, with screw caps, tared to 1 mg
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• Weighing bottles, 90 ml_, tared to 1 mg
• Weighing bottles, 90 ml_, tared to 0.1 mg
• Drying dishes, aluminum
• Timer or clock with second hand
• Electronic balance, ±0.10-mg sensitivity
• Electronic balance, ±1.0-mg sensitivity
• Watch glass, 50- and 65-mm diameters
• Evaporating dish, porcelain, 160-mm diameter, 31-mm height, with lip
• Set of 76-mm (3 in) sieves, square weave phosphor bronze wire cloth except 300 mesh
which is twilled weave. U.S. series and Tyler Screen Scale equivalent designations are as
follows:
Sand Size
vcs
cs
MS
FS
VFS
Opening
(mm)
1.0
0.5
0.25
0.105
0.047
U.S. No.
18
35
60
140
300
Tyler Mesh
Size
16
32
60
150
300
Centrifuge, International No. 11, with No. 949 rotor head, International Equip. Co., Boston,
MA
Centrifuge bottle, 500 ml_
Torsion balance
Manometer, hand-held gauge and differential pressure, PCL-200A/C Series, Omega
Engineering, Stamford, CT.
Gelatin capsules, 5 ml_
Threaded weighing bottles, 90 mL
Machined PVC caps for threaded 90-mL weighing bottles, 3.2-cm (1 1/4 in) diameter with
1.1-cm (7/16 in) diameter hole drilled in center, O-ring seal
O-rings, 3.2 x 38.1 mm (1/8 x 1 1/2 in)
Septa, rubber, 7.9-mm (5/16 in) diameter. Place in machined cap.
Hypodermic needle, 25.4 mm (1 in), 23 gauge
Ultrasonic probe, 19-mm (3/4 in) horn, 20 kHz, 300 watts
5.8.5 Reagents
Reverse osmosis (RO) water, ASTM Type III grade of reagent water
Hydrogen peroxide (H2O2), 30 to 35%
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• Sodium hexametaphosphate ((NaPO3)6), reagent grade
• Sodium carbonate (Na2CO3), reagent grade
• Sodium hexametaphosphate solution. Dissolve 35.7g of (NaPO3)6 and 7.94g of Na2CO3 in
1L of RO water. See Section 5.7.6, step 7 for standardization of sodium
hexametaphosphate solution.
• Ethyl alcohol
• Calcium sulfate (anhydrous) or equivalent desiccant
• Hydrochloric acid (HCI), 6 N, technical grade. Dilute 1L of concentrated HCI with 1L of RO
water
• Sodium carbonate (Na2CO3) reagent. Dissolve 10.6 g Na2CO3 in RO water and make to
1L(10mgCaCO3)
• 1 N sodium acetate (NaOAc) solution, buffered to pH 5. Dissolve 680 g of NaOAc in 4 L of
RO water. Add ~ 250 ml_ of acetic acid. Make to 5-L volume with RO water.
Sodium citrate solution, 0.3 M Na3C6H5O7-2H2O (88.4 g L-1)
• Sodium bicarbonate buffer solution, 1 M NaHCO3 (84 g L-1)
• Sodium dithionite (Na2S2O4 - hydrosulphite)
Saturated NaCI solution (solubility at 20°C; 360 g L-1)
• Sodium hydroxide solution (NaOH), 0.1 N. Dissolve 4 g NaOH pellets in 1 L of RO water.
5.8.6 Procedures
1. Weigh 10g of <2-mm, air-dry soil to nearest mg on an electronic balance and place into a
numbered, tared 300-mL, fleaker. Wash and tare these fleakers once every two months. A
quality control sample is included in each batch (<24 samples).
2. Add ~ 50mL of RO water and 5 to 7.5mL of H2O2 to the soil sample at ambient temperature.
Cover the soil sample with a 50-mm watch glass. Allow initial oxidation of organic matter to
complete and then place sample on hot plate. If the froth from the reaction exceeds the
capacity of the fleaker, transfer the sample to a larger beaker.
3. Place the sample on a hot plate and heat to 90°C. Add four 5 to 7.5-mL increments of H2O2
at 30-min intervals. If oxidation is incomplete, add additional H2O2 until organic matter
oxidation is complete. Heat the sample for an additional 45 min to decompose excess H2O2.
If the reaction is violent, do one or any combination of the following: (a) add small
increments of ethyl alcohol to the sample; (b) remove the sample from the hot plate to slow
the reaction; (c) transfer sample to a 1000-mL beaker; or (d) reduce the amount of H2O2 to
sample. Record any unusual sample reactions.
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4. Place the sample on the filter rack. Add 150ml_ of RO water. Insert a filter candle, connect
to the vacuum trap assembly with tubing, and turn on the vacuum. Aspirate until liquid is
removed and only slightly dampened sample remains. Wash the sample four additional
times with ~ 150ml_ of RO water. Stir the sample with filter candle to ensure all soil particles
will be rinsed. During aspiration, it may be necessary to occasionally apply back-pressure to
filter candle and remove build-up of soil which inhibits aspiration. If the sample contains
gypsum and flocculates, then the following additional washings may be used. If the sample
contains 1 to 5% gypsum, stir the sample with a magnetic stirrer for 5 minutes and wash 5
times with « 250ml_ of RO water each time. If the sample contains >5% gypsum, place the
sample in a 1000-mL beaker and stir the sample with a magnetic stirrer for 5 minutes then
wash 5 times with « 750ml_ of RO water each time to remove soluble gypsum.
5. Place sample in oven. Dry the sample overnight at 110°C. Remove the sample from the
oven, place in a desiccator, and cool to ambient temperature.
6. Record the total weight (TW) of the sample to the nearest mg.
7. Add the exact volume of sodium hexametaphosphate solution (« 10 ml_), equivalent to
0.4408 g of sodium hexametaphosphate, to each sample. Subtract the weight of the sodium
hexametaphosphate (DW) that is contained in the extracted aliquot from the silt and clay
weights to calculate silt and clay percentages. To determine the exact volume of sodium
hexametaphosphate to add to each sample, refer to Section 5.7.6, step 9. Let stand until
sample is completely moistened by sodium hexametaphosphate. Add « 175 ml_ of RO
water.
8. A sodium hexametaphosphate standardization is performed with each new batch of solution.
Use only designated weighing bottles for standardization. Wash and tare these bottles after
each standardization. Add duplicate aliquots (8.5, 9.0, 9.3, 9.6, 10.0, 10.3, 10.6, 11.0 ml_) of
sodium hexametaphosphate solution to numbered tared, 90-mL weighing bottles. Oven-dry
aliquots overnight and record dry residue weight of sodium hexametaphosphate. Determine
the exact volume of sodium hexametaphosphate to add to each sample by regressing the
volume of sodium hexametaphosphate against the dry residue weight of sodium
hexametaphosphate and then by predicting the volume needed to dispense 0.4408g of
sodium hexametaphosphate into each sample.
9. Place the sample in a horizontal shaker set at 120 oscillations min"1 and shake for 15 hours
(overnight).
10. Remove the sample from the shaker and pour through a 300-mesh (0.047-mm) sieve
mounted on a ring stand. Finger-rubbing of sample may be required during transfer to
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speed washing of sample. Place a funnel below the sieve and a 1-L cylinder below the
funnel. Collect the silt and clay in the 1-L cylinder. Avoid using jets of water in washing the
sample. Wash and rub all particles from the fleaker into the sieve. Continue to wash until
the suspension volume in the cylinder is ~ 800ml_. Sand and some of the coarse silt remain
on the sieve. Rinse all <20-(im particles into the cylinder. Fill the cylinder to 1L and cover
with a 65-mm watch glass. Place pipe insulation around sample and blank cylinders to
prevent rapid changes in temperatures. Prepare a RO water blank to measure temperature
fluctuations. Allow the cylinder to stand overnight to equilibrate the suspension with the
room temperature. Wash the sand into an evaporation dish and dry the sand at 110°C
overnight.
11. Transfer the dried sand to a nest of sieves that has a top-to-bottom order of 1.0, 0.5, 0.25,
0.1, and 0.047 mm. Shake the sand for 3 minutes on a shaker that has 1.3-cm vertical and
lateral movements and oscillates at 500 strokes min-1. Record the weight of each separate
sand fraction (SWi) to the nearest mg. If optical analysis is requested, place the very fine
sand and fine sand fractions in gelatin capsules and the remaining sand fractions in a
labeled vial. Store capsules in the labeled vial. Wash sand dishes after every use.
12. Determine the percentage of fine silt and clay gravimetrically by removing an aliquot from
the suspension in the 1-L cylinder with a Lowy, 25-mL pipet. Periodically, gravimetrically
calibrate the delivery volume of the pipet by weighing the amount of RO water dispensed
from the pipet. Record the delivery volume (DV) and use the value to calculate the results.
Regulate the vacuum such that the pipet fills in »12 seconds. Record temperature (T1) of
blank. Mount the pipet on an adjustable pipet rack (Shaw, 1932). Stir the silt and clay
suspension with mechanical stirrer for at least 5 minutes. Place the cylinder on a stable,
vibrationless table and stir with a hand stirrer in an up-and-down motion for 30 seconds.
Timing is started upon completion of the stirring. Record the time that stirring is stopped.
For the <20-jam fraction, slowly lower the closed pipet to a 10-cm depth in the suspension,
turn on the vacuum, and withdraw an aliquot at the calculated time (Table 1). Dispense the
aliquot into a tared and numbered, 90-mL weighing bottle. Rinse the pipet twice with RO
water and dispense into the tared, weighing bottle with the aliquot. For the <2-um fraction,
pipet after a time of 4.5, 5, 5.5, or 6.5 hours. Record temperature (T2) of blank. Use the
average of T1 and T2 and adjust the pipet depth in the suspension as indicated in Table 2.
Repeat the procedure described for the <20-jam fraction. If determination of carbonate is
required, use weighing bottle with screw threads. Dry the aliquots at 110°C overnight and
cool in a desiccator that contains calcium sulfate or an equivalent desiccant. Record the
weight of the residue (RW) to the nearest 0.1 mg.
13. Use the 90-mL, round-bottomed, weighing bottles for the <20-|im aliquots.
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14. Wash and tare after every fourth use. Use the 90-mL, square-bottomed, weighing bottles
for the <2-|im aliquots. Wash and tare after every use.
15. If optical mineralogy, fine-clay and/or carbonate clay determinations are not requested, the
procedural aspects of the standard air-dry PSDA for the <2-mm fraction are complete. If
optical mineralogy is requested, save the sediment and proceed on with the section on
optical mineralogy. If fine-clay and/or carbonate clay are requested, proceed on with the
sections on these analyses.
Optical Mineralogy
16. If optical mineralogy is requested, decant the suspension and transfer the sediment to a
400-mL beaker. Fill the beaker to a 5.5-cm height. Stir the sediment and allow it to settle
for 5 minutes. Discard the supernatant. Refill the beaker to 5.5-cm height. Stir again, allow
it to settle for 3 minutes, and then decant. Repeat the filling and the stirring; allow it to settle
for 2 minutes; decant until top half of suspension is clear. Transfer the sediment, which is
dominantly 20 to 50|im, to a labeled drying dish. Wash with ethanol, air-dry, and save in the
drying dish for optical mineralogy.
Fine Clay Determination (<0.2 |jm)
17. Stir the silt and clay suspension with mechanical stirrer for 5 minutes. Remove sample from
mechanical stirrer and place on table. Stir with the hand stirrer in an up-and-down motion
for 30 seconds and allow the suspension to settle for 15 minutes.
18. Pour the suspension into a centrifuge bottle and fill to the line marked on the bottle. The
marked line on each bottle is 13cm which is the distance from the center of rotation to the
surface of the suspension. Stopper and shake well to mix the suspension.
19. Balance opposite centrifuge loads, which consist of centrifuge bottle, trunnion carrier and
bucket. Place loads on a torsion balance and add water to the lighter bucket until both loads
weigh the same.
20. Read the temperature of the suspension.
21. Centrifuge at 1500 rpm.
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22. Vary the centrifuge time according to the temperature as follows:
where:
Temperature
(°C)
18
19
20
21
22
23
24
25
26
27
28
29
30
Viscosity
TI
0.01055
0.01029
0.01004
0.00980
0.00957
0.00934
0.00913
0.00892
0.00872
0.00853
0.00834
0.00816
0.00799
Delta-Density
Ap
1.501
1.501
1.502
1.502
1.502
1.502
1.502
1.503
1.503
1.503
1.503
1.504
1.504
Time
(Minutes)
39.0
38.0
37.1
36.2
35.3
34.5
33.7
32.9
32.2
31.4
30.8
30.1
29.4
= 15 cm
= 18cm
= 1 500 rpm
= 2.5 g cc-1
s
r
Nm
PP
23. After centrifuging, lower the pipet to a 3-cm depth in the suspension. Withdraw a 25-mL
aliquot at a rate of» 12 seconds. Avoid turbulence. Transfer the aliquot to a weighing
bottle.
24. Place weighing bottle with aliquot in oven. Dry overnight at 110°C. Remove sample from
oven, place in desiccator with calcium sulfate or equivalent desiccant, and cool to ambient
temperature.
25. Weigh residue weight (RW) to nearest 0.1 mg.
26. Use the 90-mL, round-bottomed, weighing bottles for the <0.2um aliquots. Wash and tare
after every fourth use.
Carbonate Clay (<2 urn): Manometer Calibration
27. Calibrate the manometer quarterly or whenever equipment changes. Calibrate by placing
replicated aliquots of 0.0 to 20.0 ml_ of the Na2CO3 reagent into numbered, tared, 90-mL
weighing bottles. Dry the standard samples in the oven overnight at 110°C. Remove
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samples from oven, place in desiccator and cool to ambient temperature. Record the weight
of the standard samples to nearest 0.1 mg.
28. Lubricate the lip of the 90-mL, weighing bottle that contains the Na2CO3 with a thin film of
glycerine. Dispense 3 ml_ of 6 N HCI into a gelatin capsule and place the top on the
capsule. If HCI leaks from the capsule, discard the capsule. Place the capsule into the
glass bottle and immediately cap the bottle. Release pressure in the bottle by piercing the
septa with a hypodermic needle which is not connected to the manometer. Allow 3 to 5
seconds for internal pressure in bottle to equalize.
29. After the gelatin capsule has dissolved (several minutes), slowly tip the bottle and rotate it to
saturate the standard sample adhering to the sides of the bottle. Avoid changing the
temperature of the container by only handling the cap. Allow sample to stand for at least 30
minutes.
30. Adjust the manometer to zero before taking measurements. Insert the hypodermic needle in
the septa stopper which is connected to the transducer. Measure the pressure inside the
weighing bottle. Record the manometer readings (mm Hg) to the nearest whole number.
31. Calculate the linear regression equation, i.e., the dependent variable is the Na2CO3 weights
(regressed or predicted values) and the independent variable is the corresponding
manometer readings.
Carbonate Clay: Analysis
32. Determine the presence of carbonates in <2-mm soil by placing soil on a spot plate and
adding two or three drops of 1 N HCI. The rate of CO2 evolution indicates the relative
amount of carbonates (1B1 b2b5).
33. If the soil contains more than a "slight" amount of carbonates, determine the amount of
carbonate clay in the <2-um dry residue. Use the 90-mL, square-bottomed, weighing bottles
for the <2-um aliquots and carbonate determination. Wash and tare after every use.
34. With a thin film of glycerine, lubricate the lip of the 90-mL, weighing bottle that contains the
<2-um residue. In each analysis batch, include an empty weighing bottle as a blank.
Dispense 3 mL of 6 N HCI into a gelatin capsule and place the top on the capsule. If HCI
leaks from the capsule, discard the capsule. Place the capsule into the glass bottle and
immediately cap the bottle. Release any pressure in the bottle by piercing the septa with a
hypodermic needle that is not connected to the manometer. Approximately 3 to 5 seconds
are required to equalize the internal pressure of the bottle.
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35. After the gelatin capsule has dissolved (several minutes), slowly tip the bottle and rotate it to
saturate the clay adhering to the sides of the bottle. Handle only the cap to avoid changing
the temperature of the container. Allow sample to stand for at least 30 minutes.
36. Adjust the manometer to zero before taking measurements. Insert the hypodermic needle in
septa stopper which is connected to the transducer. Measure the pressure inside the
weighing bottle and record the manometer readings (MR) to the nearest whole number (mm
Hg). Begin readings with the blank (BR).
37. Compare the sample readings with those of a standard curve prepared by measuring CO2
evolved from a series of Na2CO3 aliquots with a range 0 to 200 mg.
NOTE: Additional non-routine PSA method instructions described in SSIR-42.
5.8.7 Calculations
Use these calculations for all PSDA procedures. Calculations 1-6 are as follows:
1. Clay % = 100 x ((RW 2 - DW) x (CF/TW))
where:
RW2 = Residue weight (g), <2-|im fraction
DW = Dispersing agent weight (g) = (0.4408/CF)
CF =1000ml_/DV
DV = Dispensed pipet volume
TW = Total weight (g), H2O2-treated, oven-dry sample
2. Fine Silt % = 100 x ((RW20-DW) x (CF/TW)) - Clay %
where:
RW20 = Residue weight (g) of <20-jam fraction
3. Sand % = I (SWi/TW) x 100
where:
SW, = Weight of sand fractions (1.0, 0.5, 0.25, 0.1, and 0.047 mm)
4. Coarse silt % = 100 - (Clay % + Fine Silt % + Sand %)
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5. Fine Clay (%) = 100 x ((RW-DW) x (CF/TW))
where:
RW = Residue weight (g) of <0.2-um fraction
DW = Dispersing agent weight (g) = (0.4364/CF)
CF = 1000ml_/DV
DV = Dispensed pipet volume
TW = Total weight of H2O2-treated, oven-dry sample
6. Calculate carbonate clay percentage as follows:
Correct the manometer reading as follows:
CR = (MR - BR)
where:
CR = Corrected reading
MR = Manometer reading
BR = Blank reading
Three blanks are run with each batch (<24 samples). The average of three blanks is used
as BR.
Calculate two regression equations, i.e., one for corrected manometer readings <100 and
another for corrected readings >100. Use the Na2CO3 weights as the dependent variable
(regressed or predicted values) and the corresponding manometer readings as the
independent variable.
Use the corrected (CR) linear regression equations to estimate the g of CaCO3 in the
sample.
Carbonate Clay Equivalent (<2 urn) (%) = ((g CaCO3) x 100 x CF)/TW
where:
CF = 1000 mL/dispensed pipet volume (ml_)
TW = Total weight of H2O2-treated oven-dry sample
Noncarbonate Clay (<2 urn) (%) = Total Clay (%) - Carbonate Clay Equivalent (%)
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5.8.8 Report
Report each particle-size fraction to the nearest 0.1 percent.
5.8.9 Precision and Accuracy
Precision and accuracy data are available from the SSL upon request.
5.8.10 References
Bouyoucos, G.B. 1929. The ultimate natural structure of soil. Soil Sci. 28:27-37.
Drosdoff, M., and E.F. Miles. 1938. Action of hydrogen peroxide on weathered mica. Soil Sci.
46:391-395.
El Swaify, S.A. 1980. Physical and mechanical properties of Oxisols. p. 303-324. In B.K.G.
Theng (ed.) Soils with variable charge. N.Z. Soc. Soil Sci., Lower Hutt, N.Z.
Espinoza, W., R.H. Rust, and R.S. Adams Jr. Characterization of mineral forms in Andepts from
Chile. Soil Sci. Soc. Am. Proc. 39:556-561.
Gee, G.W., and J.W. Bauder. 1986. Particle-size analysis, p. 383-411. In A. Klute (ed.) Method
of soil analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9.
ASA and SSSA, Madison, Wl.
Gee, G.W., and D. Or. 2002. Particle-size analysis, p. 255-293. In J.H. Dane and G.C. Topp
(eds.) Methods of soil analysis. Part 4. Physical methods. Soil Sci. Am. Book Series No.
5. ASA and SSSA, Madison, Wl.
Grossman, R.B., and J.L. Millet. 1961. Carbonate removal from soils by a modification of the
acetate buffer method. Soil Sci. Soc. Am. Proc. 25:325-326.
Jackson, M.L. 1969. Soil chemical analysis—advanced course. 2nd ed. Univ. Wisconsin,
Madison.
Kilmer, V.J., and L.T. Alexander. 1949. Methods of making mechanical analyses of soils. Soil
Sci. 68:15-24.
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Kilmer, V.J., and J.K. Mullins. 1954. Improved stirring and pipeting apparatus for mechanical
analysis of soils. Soil Sci. 77:437-441.
Kubota, T. 1972. Aggregate formation of allophonic soils; Effects of drying on the dispersion of
soils. Soil Sci. and Plant Nutr. 18:79-87.
Maeda, T., H. Takenaka, and B.P. Warkentin. 1977. p. 229-263. Physical properties of
allophone soils. In N.C. Brady (ed.) Adv. Agron. Acad. Press, Inc., New York, NY.
Mehra, O.P., and M.L. Jackson. 1960. Iron oxide removal from soils and clays by a dithionite-
citrate system buffered with sodium bicarbonate, p. 237-317. In Clays and clay minerals.
Proc. 7th Conf. Natl. Acad. Sci. Natl. Res. Counc. Pub., Washington, DC.
Mikhail, E.H., and G.P. Briner. 1978. Routine particle size analysis of soils using sodium
hypochlorite and ultrasonic dispersion. Aust. J. Soil Res. 16:241-244.
Rabenhorst, M.C., and L.P. Wilding. 1984. Rapid method to obtain carbonate-free residues from
limestone and petrocalcic materials. Soil Sci. Soc. Am. J. 48:216-219.
Saly, R. 1967. Use of ultrasonic vibration for dispersing of soil samples. Sov. Soil Sci. 11:1547-
1559.
Shaw, T.M. 1932. New aliquot and filter devices for analytical laboratories. Ind. Eng. Chem.
Anal. Ed. 4:409.
Shields, L.G., and M.W. Meyer. 1964. Carbonate clay: Measurement and relationship to clay
distribution and cation exchange capacity. Soil Sci. Soc. Am. Proc. 28:416-419.
Skopp, J. 1992. Concepts of soil physics. University of Nebraska, Lincoln, NE.
Soil Survey Division Staff. 1993. Soil survey manual. USDA. Handb. No. 18. U.S. Govt. Print.
Office, Washington, DC.
Soil Survey Staff. 1996. Soil survey laboratory methods manual. Version No. 3.0. USDA-NRCS.
Soil Survey Investigations Report No. 42. U.S. Govt. Print. Office, Washington, DC.
Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys. 2nd ed. Agric. Handb. No. 436. USDA-NRCS. U.S. Govt. Print.
Office, Washington, DC.
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Sumner, M.E., 1992. The electrical double layer and clay dispersion, p. 1-32. In M.E. Sumner
and B.A. Stewart (eds.) Soil crusting, chemical, and physical processes. CRC Press Inc.,
Boca Raton, FL
Watson, J.R. 1971. Ultrasonic vibration as a method of soil dispersion. Soil Fertil. 34:127-134.
Williams, D.E. 1948. A rapid manometric method for the determination of carbonate in soils. Soil
Sci. Soc. Am. Proc. 13:127-129.
5.9 Water Retention, 1500-kPa
5.9.1 Summary of Method
The pressure desorption procedure (U.S. Salinity Laboratory Staff, 1954) is used. A sample of
<2-mm (sieved), air-dry soil is placed in a retainer ring sitting on a cellulose membrane in a
pressure-membrane extractor. The membrane is covered with water to wet the samples by
capillarity. The sample is equilibrated at 1500 kPa. The pressure is kept constant until
equilibrium is obtained (Klute, 1986). The gravimetric water content is determined.
5.9.2 Interferences
A leaking pressure extractor prevents equilibration of samples. Check outflow air to verify that
the pressure membrane extractor is functioning properly and does not leak. The pressure
should be monitored for stability. Equilibration must be done at constant temperature and
humidity. Samples that do not wet by capillarity are moistened with ethyl alcohol.
Laboratory-determined, water retention data are usually higher than field-determined, water
retention data because the confining soil pressure is not present in the laboratory (Bruce and
Luxmoore, 1986).
Aerated 0.005 M CaSO4 has also been recommended (Dane and Hopmans, 2002), especially
for fine-textured soils that contain significant amounts of swelling clays. Distilled or deionized
water can possibly promote dispersion of clays in samples and freshly drawn tap water is often
supersaturated with air, affecting the water content at a given pressure head (Dane and
Hopmans, 2002).
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5.9.3 Safety
High-pressure plumbing must be maintained in good working order. Ensure that the pressure is
zero before removing bolts from the pressure-apparatus lid. Ensure that the bolts are tightened
before applying pressure. Do not drop the heavy lid.
5.9.4 Equipment
• Pressure membrane extractor (Fig.1 and 2)
• Cellulose membrane
• Retainer rings. Use 10-mm height and 50-mm diameter rings for organic soils and 10-mm
height and 40-mm diameter rings for all other soils.
• Electronic balance, ±0.01-g sensitivity
Oven, 110°C
• Pressure source, regulator, and gauge
• Metal weighing cans, tared, with lids
• Vacuum trap assembly
Vacuum, 80 kPa (0.8 bar)
5.9.5 Reagents
• Ethyl alcohol, 95%, technical grade
• Reverse osmosis (RO) water
5.9.6 Procedure
1. Submerge a cellulose membrane in RO water for 12 h or more before use. Install the wet
cellulose membrane in the pressure extractor.
2. Add water and retaining rings. Add enough water to keep membrane moist. Water level
should be less than height of retaining rings. Use 5-cm diameter rings for soils that are
>12% in organic matter. Use 4-cm diameter rings for all other soils.
3. Fill retaining rings with 10 to 15 g of <2-mm or fine-grind, air-dry soil sample. Include a
quality control (QC) sample with each plate. Continue to add water until all samples have
moistened by capillarity. If samples do not moisten, apply ethyl alcohol to the surface of the
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sample. Allow ethyl alcohol to evaporate. Cover samples with a sheet of plastic to reduce
evaporation, close the extractor, and let stand overnight.
4. Remove excess water on the plate with a vacuum and trap assembly.
5. Assemble the extractor and uniformly tighten the bolts. Torque the bolts on both sides of
the hinge to 138.0 kPa (200 psi). Torque the remaining bolts to 103.5 kPa (150 psi).
6. Increase air pressure « 150 kPa every 15 min until 1500 kPa is reached. After 4 h, apply the
pressure differential by closing the valve that joins the mercury circuit and by opening the
pressure release valve until air is forced through the mercury. Quickly close the pressure
release valve. This forces the rubber diaphragm against the top of the samples. The
samples are equilibrated when water ceases to emit from the outflow tube.
7. At equilibrium, open the extractor and quickly transfer the samples to water cans, cover with
lids, and record the weights (Ms+w).
8. Remove the lids, place samples in the oven, and dry at 110°C overnight. Remove samples
from the oven, replace the lids, allow cans to cool to ambient temperature, and record the
weights (Ms).
9. Record the weights of the empty cans (Me).
5.9.7 Calculations
H20 % = 100 x [(Ms+w - Ms)/ (Ms - Mc)]
where:
H2O % = Percent gravimetric water content
MS+W = Weight of solids + H2O + container
MS = Weight of solids + can
Mc = Weight of container
Gypsiferous soils are a special case because gypsum (CaSO4-2H2O) loses most of its two water
molecules at 105°C. Properties of gypsiferous soils such as 1500-kPa water content that are
reported on an oven-dry weight basis are converted to include the weight of crystal water in
gypsum. Refer to procedure 3D3 for these conversion calculations. The 1500-kPa water
content is corrected when the gypsum content of the soil is >1 %. Gypsum content of the soil is
determined in procedure 4E2a1a1.
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5.9.8 Report
Report water content to the nearest 0.1 percent.
5.9.9 Precision and Accuracy
Precision and accuracy data are available from the SSL upon request.
5.9.10 References
Bruce, R.R., and R.J. Luxmoore. 1986. Water retention: Field methods, p. 663-686. In A.
Klute (ed.) Methods of soil analysis. Part 1. Physical and mineralogical methods. 2nd
ed. Agron. Monogr. 9. ASA and SSSA, Madison, Wl.
Dane, J.H., and J. W. Hopmans. 2002. Water retention and storage: Laboratory, p. 675-720.
In J.H. Dane and G.C. Topp (eds.) Methods of soil analysis. Part 4. Physical methods.
Soil Sci. Am. Book Series No. 5. ASA and SSSA, Madison, Wl.
Klute, A. 1986. Water retention: Laboratory methods, p. 635-662. In A. Klute (ed.) Methods of
soil analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9.
ASA and SSSA, Madison, Wl.
Nelson, R.E., L.C. Klameth, and W.D. Nettleton. 1978. Determining soil content and
expressing properties of gypsiferous soils. Soil Sci. Soc. Am. J. 42:659-661.
U.S. Salinity Laboratory Staff. 1954. L.A. Richards (ed.) Diagnosis and improvement of saline
and alkali soils. USDA Agric. Handb. 60. U.S. Govt. Print. Office, Washington, DC.
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5.10 Calcium Carbonate Equivalent
5.10.1 Summary of Method
The amount of carbonate in the soil is measured by treating the samples with HCI. The evolved
CO2 is measured manometrically. The amount of carbonate is then calculated as percent
CaCO3.
5.10.2 Interferences
Chemical interference is the reaction by the acid with other carbonates, e.g., carbonates of Mg,
Na, and K, that may be present in soil sample. The calculated CaCO3 is only a semiquantitative
measurement (Nelson, 1982). Analytical interference may be caused by temperature changes
within the reaction vessel. When sealing the vessel, the analyst should not hold the vessel any
longer than necessary to tighten the cap. The internal pressure must be equalized with the
atmosphere. After the septum has been pierced with a needle, « 5 to 10 s are required to
equalize the internal pressure of the bottle. With extensive use, the septa leak gas under
pressure. The septa should be replaced at regular intervals. The analyst should not touch the
glass of the vessel when reading the pressure.
5.10.3 Precautions
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses) when handling acids. Thoroughly wash hands after
handling acids. Use the fume hood when diluting concentrated HCI. Use the safety showers
and eyewash stations to dilute spilled acids. Use sodium bicarbonate and water to neutralize
and dilute spilled acids.
The gelatin capsule may leak acid while being filled. Keep other personnel away from the area
when filling capsules.
High pressure may develop inside the bottle if there is a large amount of calcareous sample.
Do not use more than 2 g of any sample in the bottles. If high pressure develops in the bottle,
release the pressure by venting the gas with a syringe needle. Some bottles may break without
shattering. Discard any bottle with hairline cracks or obvious defects.
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5.10.4 Equipment/Materials
• Electronic balance, ±0.10-mg sensitivity
• Electronic balance, ±1mg-sensitivity
• Threaded weighing bottles, wide-mouth, clear glass, standard, 120 ml_ (4 fl. oz.), 48-mm
neck size. For best results, grind rim of bottle with 400-600 grit sandpaper on a flat glass
plate.
• Machined PVC caps for threaded 120-mL (4 fl. oz.) weighing bottles, 54-mm diameter with
12.7-mm diameter hole drilled in center, O-ring seal.
O-rings, 3.2 x 50.8 x 57.2 mm (1/8 x 2 x 2 1/4 in)
• Flanged stopper No. 03-255-5, Fisher Scientific. Place in machined cap.
• Manometer, hand-held gauge and differential pressure, PCL-200 Series, Omega
Engineering, Stanford, CT.
• Hypodermic needle, 25.4 mm (1 in), 23 gauge. Connect needle to pressure tubing on
transducer.
• Mechanical rotating shaker, 140 rpm, Eberbach 6140, Eberbach Corp., Ann Arbor, Ml.
5.10.5 Reagents
• Reverse osmosis deionized (RODI) water, ASTM Type I grade of reagent water
• Methyl red indicator
• Hydrochloric acid (HCI), concentrated, 12 N
HCI, 3 N. Dilute 500 ml_ of concentrated HCI with 1500 ml_ RODI water. Add a few
crystals of methyl red indicator. Methyl red indicator will turn yellow if HCI is consumed by
sample. If this reaction occurs, adjust the sample size (smaller).
• Gelatin capsule, 10 ml_, size 11, Torpac Inc., Fairfield, NJ.
• Glycerin, USP. Put the glycerin in a small squeeze bottle and use as a lubricant for the O-
rings.
• CaCO3, Ultrex, assay dried basis 100.01%.
5.10.6 Procedure
Manometer Calibration: Calibrate the manometer quarterly or whenever equipment changes
(e.g., old rubber septum replaced). Calibrate by weighing three replicates of CaCO3 standards
(0, 0.025, 0.05, 0.1, 0.2, 0.3, 0.4, 0.5, 0.75 g). Weigh to the nearest 0.1 mg. Dry the standard
samples in the oven for 2 h at 110°C. Remove samples from oven, place in desiccator and cool
to ambient temperature.
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<2-mm Basis: A CaCI2 pH >6.95 is generally used as an indicator of the presence of
carbonates. The presence of carbonates (effervescence with HCI) is also checked during lab
preparation.
1. Weigh 0.5 to 2 g of fine-grind, air-dry soil sample to the nearest mg and place in a 120-mL,
wide-mouth bottle. Run 3 blanks and a quality control check sample with every batch of 24
samples. The quality control check sample serves as a single point check. Vary the sample
weight according to the CaCO3 content based on effervescence of sample as follows:
2. Use a 2-g sample weight if effervescence is None, Very Slight, or Slight.
3. Use a 1-g sample weight, if effervescence is Strong.
4. Use a 0.5-g sample weight, if effervescence is Violent.
5. Lubricate the O-ring of bottle cap with glycerin from a squeeze bottle.
6. Dispense 10 mL of 3 N HCI into a gelatin capsule and carefully place the top on the capsule.
The HCI may squirt or leak out of capsule. If this happens, discard the capsule.
7. Place the capsule in bottle and cap bottle immediately.
8. Release any pressure in the bottle by piercing the stopper with a hypodermic needle.
Remove the needle after« 5 to 10 s.
9. After 5 to 10 min, the HCI dissolves through the capsule. Shake the bottle at a rate of 140
rpm on the shaker for the first 10 min and last 10 min of a 1-h interval at room temperature
(20°C ± 2°C). After this 1 h, measure the pressure in the bottle by piercing the stopper of the
cap with a hypodermic needle connected to the manometer.
10. Auto-zero the manometer before taking readings. Record the manometer readings (mm Hg).
5.10.7 Calculation
Correct the manometer readings as follows:
CR = (MR-BR)
where:
CR = Corrected reading
MR = Manometer reading
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BR = Blank reading
Three blanks are run with each batch of 24 samples. The average of three blanks is used as
BR.
Calculate the regression equation for the corrected manometer readings. Use the CaCO3
weights as the dependent variable (regressed or predicted values) and the corresponding
manometer readings as the independent variable.
Use the corrected (CR) linear regression (slope, intercept) equation to estimate % CaCO3 in the
sample as follows:
CCE = [(CR x Slope + Intercept) /Sample Weight (g)] x AD/OD
where:
CCE = Calcium Carbonate Equivalent (%) in <2-mm fraction or 2- to 20-mm fraction
CR = Corrected manometer reading
AD/OD= Air-dry/oven-dry ratio (procedure 3D1)
Carbonate = (A x B) + [C x (1-B)]
where:
Carbonate = Carbonate as CaCO3 on a <20-mm basis (%)
A = CaCO3 in <2-mm fraction (%)
B = Weight of the <20-mm fraction minus the weight of the 2- to 20-mm fraction divided
by the weight of <20-mm fraction (procedures 1B2b2f and 3A2).
C = CaCO3 in 2- to 20-mm fraction (%)
5.10.8 Reporting
Report CaCO3 equivalent as a percentage of oven-dry soil to the nearest whole number.
5.10.9 References
Amer, F.A., A. Mahmoud, and V. Sabel. 1985. Zeta potential and surface area of calcium
carbonate as related to phosphate sorption. Soil Sci. Soc. Am. J. 49:1137-1142.
Boischot, P, M. Coppenet, and J. Hebert. 1950. Fixation de I'acide phosphorique surle calcaire
des sols. Plant Soil 2:311-322.
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Loeppert, R. H., and D. L. Suarez. 1996. Carbonate and gypsum, p. 437-474. In D.L. Sparks
(ed.) Methods of Soil Analysis. Part 3 - Chemical methods. Soil Sci. Am. Book Series
No. 5. ASA and SSSA, Madison, Wl.
Nelson, R.E. 1982. Carbonate and gypsum, p. 181-197. In A.L Page, R.H. Miller, and D.R.
Keeney (eds.) Methods of soil analysis. Part 2. Chemical and microbiological properties.
2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, Wl.
Rabenhorst, M.C., L.T. West, and L.P. Wilding. 1991. Genesis of calcic and petrocalcic horizons
in soils over carbonate rocks, p. 61-74. In W.D. Nettleton (ed.) Occurrence,
characteristics, and genesis of carbonate, gypsum, and silica accumulations in soils. Soil
Sci. Soc. Am. Spec. Publ. No. 26. ASA and SSSA, Madison, Wl.
Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys. USDA-NRCS Agric. Handb. 436. 2nd ed. U.S. Govt. Print.
Office, Washington, DC.
Talibudeen, O., and P. Arambarri. 1964. The influence of the amount and the origin of calcium
carbonates on the isotopically-exchangeable phosphate in calcareous soils. J. Agric. Sci.
62:93-97.
5.11 Total Carbon, Total Nitrogen, and Total Sulfur
5.11.1 Summary of Method
An air-dry (80 mesh, <180 urn) sample is packed in a tin foil, weighed, and analyzed for total C,
N, and S by an elemental analyzer. The elemental analyzer works according to the principle of
catalytic tube combustion in an oxygenated CO2 atmosphere and high temperature. The
combustion gases are freed from foreign gases. The desired measuring components (N2, CO2,
and SO2) are separated from each other with the help of specific adsorption columns and
determined in succession with a thermal conductivity detector, with helium as the flushing and
carrier gas.
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5.11.2 Interferences
Contamination through body grease or perspiration must be avoided in sample packing.
Substance loss after weighing should be avoided by exact folding of the sample into the tin foil.
Air in the sample material should be minimized (falsifying the N value) by compressing the
sample packing. Insufficient O2 dosing reduces the catalysts, decreasing their effectiveness
and durability. Burnt sample substance that remains in ash finger falsifies the results of
subsequent samples. WO3 is used as sample additive and combustion filling to aid combustion
or bind interfering substances (alkaline or earth-alkaline elements, avoid non-volatile sulfates).
5.11.3 Precautions
Exhaust gas pipes should lead into a ventilated fume hood. Aggressive combustible products
should not be analyzed. Before working on electrical connections (adsorption columns) or
before changing reaction tubes, the instrument must be cooled down and cooled off. Gloves
and safety glasses should be worn at all times during operation and maintenance of instrument.
5.11.4 Equipment/Materials
Elemental analyzer with on-line electronic balance (0.1 mg sensitivity) and automatic sample
feeder, ElementarvarioELand Elementar varioEL III, Elementar Analysensysteme GmbH,
Hanau-Germany, and combustibles (Elementar Americas, Inc., Mt. Laurel, NJ; Alpha Resources
Inc., Stevensville, Ml) as follows:
• Quartz ash finger, quartz
• Quartz bridge
• Combustion tube
• Reduction tube
• Gas purification (U-tube, GL 18)
• Support tube (65 mm)
• Protective tube
• O2 lance (150 mm rapid N)
• Tin boats (4x4x11 mm)
• Tin foil cups
• Computer, with varioEL software, Elementar Analysensysteme GmbH, Hanau-Germany,
and printer.
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5.11.5 Reagents
Sulfanilic acid, calibration standard, 41.6% C, 4.1% H, 8.1 % N, 27.7% O, and 18.5% S
• Copper sticks
• Corundum balls, high purity, alumina spheres, 3-5 mm
• Cerium dioxide, 1-2 mm
• Tungsten oxide powder, sample additive
• Tungsten trioxide granulate, combustion tube filling
• Quartz wool
• Silver wool
• Phosphorus pentoxide, Sicapent, Elementar Americas, Inc., Mt. Laurel, NJ
• Helium, carrier gas, 99.996% purity
• Oxygen, combustion gas, 99.995% purity
5.11.6 Procedure
1. Elemental Analyzer Set-up and Operation - Refer to the manufacturer's manual for
operation and maintenance of the elemental analyzer. Conditioning of the elemental
analyzer and determination of factor and blank value limit are part of the daily measuring
routine. The following are only very general guidelines for instrument parameters for the
various analytes in the CMS mode.
a. Temperature
Furnace 1 1140°C
Furnace 2 850 ° C
Furnace 3 0 ° C
CO2 Column 85 ° C
SO2 Column 210 ° C
SO2 Col. Standby 140 ° C
b. Timing
Flush 5 s
Oxygen Delay 10s
Auto Zero Delay 30 s
Integrator Reset Delay 50 s
Peak Anticipation N 70 s
Peak Anticipation C 125 s
Peak Anticipation S 70 s
c. Integrated Reset Delay for S 60 s
d. Thresholds
N Peak 3 mV
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C Peak 3 mV
S Peak 3 mV
e. O2 Dosing
Index 1 150s
Index 2 1 s
Index 3 90s
Index 4 120s
Index 5 180s
2. Elemental Analyzer Calibration and Analysis
a. A calibration that covers the desired working range of each element is performed.
The test substances sulfanilic acid with each given element content and different
weights is analyzed. The PC program automatically computes the calibration
function (linear, polynomial, or mixed). Calibration will typically remain stable for at
least 6 months. Re-calibration is recommended when the daily factor is outside the
range of 0.9 to 1.1 or if components that influence the results (e.g., detector or
adsorption column) have been exchanged. Changing the desorption temperature of
adsorption columns can also require a re-calibration.
b. Add 0.100 g of tungsten oxide in tin foil and tare. An homogenized fine-grind, air-dry
soil sample is then packed in this tin foil, weighed (0.100 to 0.05 g), and placed into
the carousel of the automatic sample feeder of elemental analyzer. Sample weight is
based on visual observation of the sample, related to element content, homogeneity,
and combustion behavior of the sample. The sample weight is entered in the PC
from an on-line electronic balance via an interface. A quality control (QC) sample is
performed at a minimum of every 35 to 40 samples.
5.11.7 Calculation
C(%) = CiXAD/OD
where:
C (%) = C (%), oven-dry basis
Ci = C (%) instrument
AD/OD = Air dry/oven-dry ratio
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N (%) = NiX AD/OD
where:
N (%) = N (%), oven-dry basis
Ni = N (%) instrument
AD/OD = Air-dry/oven-dry ratio
S (%) = Si x AD/OD
where:
S (%) = S (%) on oven-dry basis
Si = S (%) instrument
AD/OD = Air-dry/oven-dry ratio
5.11.8 Reporting
Report total C and S percentages to the nearest 0.01% and total N to the nearest 0.001%.
5.11.9 References
Beaton, James D., G.R. Burns, and J. Platou. 1968. Determination of sulphur in soils and plant
material. Tech. Bull. No. 14. The Sulfur Inst., Washington, DC.
Bremmer, J.M. 1996. Nitrogen-Total, p. 1085-1121. In D.L Sparks (ed.) Methods of soil
analysis. Part 3. Chemical methods. No. 5. ASA and SSSA, Madison, Wl.
Bremmer, J.M., and C.S. Mulvaney. 1982. Nitrogen -Total, p. 595-624. In A.L Page, R.H.
Miller, and D.R. Keeney (eds.) Methods of soil analysis. Part 2. Chemical and
microbiological properties. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, Wl.
Broadbent, F.E. 1953. The soil organic fraction. Adv. Agron. 5:153-183.
Dumas, J.B.A. 1831. Precedes de I'analyse organique. Ann. Chim. Phys. 247:198-213.
Kjeldahl, J. 1883. Neue Methode zur Bestimmung des Stickstoffs in organischen Korpern. Z.
Anal. Chem. 22:366-382.
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Nelson, D.W., and L.E. Sommers. 1996. Total carbon, organic carbon, and organic matter, p.
961-1010. In D.L Sparks (ed.) Methods of soil analysis. Part 3. Chemical methods. No.
5. ASA and SSSA, Madison, Wl.
Nommik, H., and K. Vahtras. 1982. Retention and fixation of ammonium and ammonia in soils.
P. 123-171. In F.J. Stevenson (ed.) Nitrogen in agricultural soils. Agronomy 22, ASA and
SSSA, Madison, Wl.
Soil Science Society of America. 1987. Glossary of soil science terms. Rev. ed. Soil Sci. Soc.
Am., Madison, Wl.
Soil Survey Staff. 1996. Soil survey laboratory methods manual. Version No. 3.0. USDA-NRCS.
Soil Survey Investigations Report No. 42. U.S. Govt. Print. Office, Washington, DC.
Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys. USDA-NRCS Agric. Handb. 436. 2nd ed. U.S. Govt. Print.
Office, Washington, DC.
Tabatabai, M.A. 1996. Sulfur, p. 921-960. In D.L. Sparks (ed.) Methods of soil analysis. Part 3.
Chemical methods. No. 5. ASA and SSSA, Madison, Wl.
Young, J.L., and R.W. Aldag. 1982. Inorganic forms of nitrogen in soil. p. 43-66. In F.J.
Stevenson (ed.) Nitrogen in agricultural soils. Agronomy 22. ASA and SSSA, Madison,
Wl.
5.12 pH
5.12.1 Summary of Method
The pH is measured in soil-water (1:1) and soil-salt (1:2 CaCI2) solutions. For convenience, the
pH is initially measured in water and then measured in CaCI2. With the addition of an equal
volume of 0.02 M CaCI2 to the soil suspension that was prepared for the water pH, the final soil-
solution ratio is 1:2 0.01 M CaCI2.
A 20-g soil sample is mixed with 20 ml_ of reverse osmosis (RO) water (1:1 w:v) with occasional
stirring. The sample is allowed to stand 1 h with occasional stirring. The sample is stirred for 30
s, and the 1:1 water pH is measured. The 0.02 M CaCI2 (20ml_) is added to soil suspension, the
sample is stirred, and the 1:2 0.01 M CaCI2 pH is measured (4C1a2a2).
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5.12.2 Interferences
The pH will vary between the supernatant and soil sediment (McLean, 1982). Measure the pH
just above the soil sediment to maintain uniformity. Clays may clog the KCI junction and slow
the electrode response. Clean the electrode. Wiping the electrode dry with cloth, laboratory
tissue or similar material may cause electrode polarization. Rinse the electrode with distilled
water and pat dry.
Atmospheric CO2 affects the pH of the soil:water mixture. Closed containers and nonporous
materials will not allow equilibration with CO2. At the time of pH determination, the partial
pressure of CO2 and the equilibrium point must be considered, if doing critical work.
5.12.3 Precautions
No significant hazards are associated with the procedure. Follow standard laboratory safety
practices.
5.12.4 Equipment/Materials
• Measuring scoop, handmade, ~ 20-g capability
• Paper cup, 120 ml_ (4 fl. oz.), disposable, Solo Cup Co., No. 404
• Dispenser, 0 to 30 ml_, Repipet or equivalent
• Beverage stirring sticks, wood
• Titration beakers, polyethylene, 250 ml_
• Automatic titrator, Metrohm Titroprocessors, Control Units, Sample Changers, and
Dosimats, Metrohm Ltd., Brinkmann Instruments, Inc.
• Combination pH-reference electrode, Metrohm part no. 6.0210.100, Brinkmann
Instruments, Inc.
5.12.5 Reagents
• Reverse osmosis (RO) water, ASTM Type III grade of reagent water
• Borax pH buffers, pH 4.00, pH 7.00, and pH 9.18, for electrode calibration, Beckman,
Fullerton, CA.
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• Calcium chloride (CaCI2), 0.02 M. Dissolve 23.52 g of CaCI2'2H2O in RO water and dilute
to8L
5.12.6 Procedure
1. Use a calibrated scoop to measure « 20 g of <2-mm or fine-grind, air-dry soil. If sample is
moist, use calibrated scoop to achieve « 20 g of air-dry soil.
2. Place the sample in a 120-mL (4-oz) paper cup.
3. Dispense 20 ml_ of RO water into sample and stir.
4. Place paper cup with sample in 250-mL titration beaker, allow to stand for 1 h, stirring
occasionally.
5. Load beakers into sample changer.
6. Calibrate the pH meter using the pH 9.18, 7.00 and pH 4.00 buffer solutions.
7. Sample stirring, waiting interval for readings, addition of CaCI2 solution, pH readings, and
rinsing of electrode are controlled by computer.
8. The general sequence used by the automated system is as follows:
o The sample is lifted so that the pH electrode is positioned above the soil
sediment.
o The sample is stirred for 30 s.
o After 1 min, 1:1 water pH is read. Record pH to the nearest 0.01 unit.
o The 20 ml_ of 0.02 M CaCI2 are added to sample. The sample is stirred for 30 s.
o After 1 min, the 1:2 CaCI2 pH is read. Record pH to the nearest 0.01 unit.
o The sample is lowered, and the electrode and stirrer are rinsed with RO water.
o The next sample is positioned for analysis.
o The cycle is repeated until all samples have been analyzed.
5.12.7 Calculation
No calculations are required for this procedure.
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5.12.8 Reporting
Report the 1:1 water pH and the 1:20.01 MCaCI2 pH to the nearest 0.1 pH unit.
5.12.9 References
Foth, H.D., and E.G. Ellis. 1988. Soil fertility. John Wiley and Sons. NY, NY. McLean, E.G.
1982. Soil pH and lime requirement, p. 199-224. In A.L Page, R.H. Miller, and D.R.
Keeney (eds.) Methods of soil analysis. Part 2. Chemical and microbiological properties.
2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, Wl.
Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys, USDA-NRCS Agric. Handb. 436. 2nd ed. U.S. Govt. Print.
Office, Washington, DC.
5.13 BaCI2-Triethanolamine, pH 8.2 Extraction
5.13.1 Summary of Method
A soil sample is leached with a BaCI2-TEA solution buffered at pH 8.2. Sample is allowed to
stand overnight, shaken, and centrifuged. The extract is back-titrated with HCI. The difference
between a blank and the extract is the extractable acidity. Extractable acidity is reported in meq
100 g"1 soil or (cmol (+) kg"1).
5.13.2 Interferences
No significant interferences are known to exist with this method. However, for some very acid
soils, the buffer capacity of the BaCI2-TEA solution may be exceeded.
5.13.3 Safety
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses) when preparing reagents, especially concentrated acids and
bases. Dispense concentrated acids in a fume hood. Thoroughly wash hands after handling
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reagents. Use the safety showers and eyewash stations to dilute spilled acids and bases. Use
sodium bicarbonate and water to neutralize and dilute spilled acids.
5.13.4 Equipment
Electronic balance, ±1.0-mg sensitivity
Pipettes or dispenser, adjustable volume to 40 mL
Vortexer, mini, Analog, VRW Scientific Products
Centrifuge tubes, 50-mL, polyethylene
Centrifuge, Centra, GP-8, Thermo IEC, Needham Heights, MA
Titration beakers, 250-mL, plastic, Metrohm Ltd., Brinkmann Instruments Inc.
Automatic titrator, with control unit, sample changer, and dispenser, Metrohm Ltd.,
Brinkmann Instruments, Inc.
Combination pH-reference electrode, Metrohm Ltd., Brinkmann Instruments, Inc.
Computer, with Titrino Workcell software, Metrohom Ltd., Brinkmann Instruments, Inc., and
printer
5.13.5 Reagents
Reverse osmosis deionized (RODI) water
Hydrochloric acid (HCI), concentrated, 12 N
HCI, 0.13 N, standardized. Dilute 193 mL of concentrated HCI to 16-L volume with RODI
water.
Buffer solution (0.5 N BaCI2, 0.2 N Triethanolamine (TEA), pH 8.2). Dissolve 977 g of
BaCI2-2H2O in 8 L of RODI water. Dissolve 477 g of TEA in 4 L of RODI water. Mix two
solutions and bring to nearly 16-L volume with RODI water. Adjust to pH 8.2 with « 33 mL
of concentrated HCI or barium hydroxide. Bring to 16-L volume with RODI water.
Replacement solution. Dissolve 977 g of BaCI2-2H2O in 8 L of RODI water. Add 80 mL of
buffer solution and dilute to 16-L volume with RODI water.
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5.13.6 Procedure
Extraction of Acidity
1. Weigh 5 g of <2-mm or fine-grind, air-dry soil to the nearest mg and place in a centrifuge
tube. If sample is moist, weigh enough soil to achieve ~ 5 g of air-dry soil. Prepare at least
two reagent blanks (no sample in tube) and one quality control check sample per 21
samples.
2. Add 40.00 ml_ of BaCI2-TEA solution to sample. Cap the tube and shake to ensure all soil is
wetted. Place tube in a rack.
3. Place tube rack on its side and gently shake to stratify the mixture lengthwise along the
tube. Allow to stand overnight on its side.
4. Centrifuge sample at 2000 rpm for 5 min.
5. Decant extract into numbered titration beakers.
6. Add 40 ml_ of replacement solution to sample.
7. Cap tube and use a vortexer to loosen soil. Manually shake.
8. Repeat Sections 7.4 - 7.7.
9. Repeat Sections 7.4 - 7.5. Total volume in titration beaker should be ~ 120 ml_.
Titration of BaCI2-TEA Extract
10. Place titration beakers on automatic sample changer.
11. Refer to the manufacturer's manual for operation of the automatic titrator.
12. Calibrate the titrator meter with 9.18, 7.00 and 4.00 pH buffers. Set-up the automatic titrator
to sent end point mode. The "Set" pH parameters are listed as follows:
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Parameter Value
Ep1 pH 4.60
Dyn change pH 1.5 units
Drift 0.4 mV s-1
Time delay 10s
Drift 0.4 mV s-1
Temp 25°C
Stop Volume 75 ml_
13. If pre-titration pH is 0.3 units lower than the average pH of the blanks, re-run using a 0.5-g
sample.
14. Record the titer to the nearest 0.01 ml_. Record the normality of the HCI solution. Average
the titer of the reagent blanks and record.
5.13.7 Calculations
Extractable acidity (meq 100 g'1) = {[(B - T) x N x R]/C} x 100
where:
B = Average reagent blank titer (ml_)
T = Sample titer (ml_)
N = Normality of HCI
C = Sample Weight (g)
100 = Conversion factor (100-g basis)
R = Air-dry/oven-dry ratio (procedure 3D1) or field-moist/oven-dry ratio
(procedure 3D2)
5.13.8 Report
Report extractable acidity to the nearest 0.1 meq 100 g"1 (cmol (+) kg"1).
5.13.9 Precision and Accuracy
Precision and accuracy data are available from the SSL upon request.
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5.13.10 References
Holmgren, G.G.S., R.L. Juve, and R.C. Geschwender. 1977. A mechanically controlled variable
rate leaching device. Soil Sci. Am. J. 41:1207-1208.
5.14 Cation Exchange Capacity (Total)
5.14.1 Summary of Method
Displacement after washing is the basis for this procedure. The CEC is determined by
saturating the exchange sites with an index cation (NH4+); washing the soil free of excess
saturated salt; displacing the index cation (NH4+) adsorbed by the soil; and measuring the
amount of the index cation (NH4+). A sample is leached using 1 N NH4OAc and a mechanical
vacuum extractor (Holmgren et al., 1977). The extract is weighed and saved for analyses of the
cations. The NH4+ saturated soil is rinsed with ethanol to remove the NH4+ that was not
adsorbed. The soil is then rinsed with 2 M KCI. This leachate is then analyzed by steam
distillation and titration to determine the NH4+ adsorbed on the soil exchange complex. The
CEC by NH4OAc, pH 7 is reported in meq 100 g"1 or (cmol (+) kg"1) soil in procedure
4B1a1a1a1.
5.14.2 Interferences
Incomplete saturation of the soil with NH4+ and insufficient removal of NH4+ are the greatest
interferences to this method. Ethanol removes some adsorbed NH4+ from the exchange sites of
some soils. Isopropanol rinses have been used for some soils in which ethanol removes
adsorbed NH4+. Soils that contain large amounts of vermiculite can irreversibly "fix" NH4+. Soils
that contain large amounts of soluble carbonates can change the extractant pH and/or can
contribute to erroneously high cation levels in the extract. This method overestimates the "field"
CEC of soils with pH <7 (Summer and Miller, 1996).
5.14.3 Safety
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses) when preparing reagents, especially concentrated acids and
bases. Dispense concentrated acids and bases in a fume hood. Thoroughly wash hands after
handling reagents. Use the safety showers and eyewash stations to dilute spilled acids and
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bases. Use sodium bicarbonate and water to neutralize and dilute spilled acids. Nessler's
reagent contains mercury which is toxic. Proper disposal of the Nessler's reagent and clean-up
of equipment in contact with the reagent is necessary.
Ethanol is flammable. Avoid open flames and sparks. Standard laboratory equipment includes
fire blankets and extinguishers for use when necessary. Follow the manufacturer's safety
precautions when using the vacuum extractor and the Kjeltec Auto Analyzers.
5.14.4 Equipment
Electronic balance, ±1.0-mg sensitivity
Mechanical vacuum extractor, 24-place, SAMPLETEX, MAVCO Industries, Lincoln, NE
(Fig. 1 and 2)
Tubes, 60-mL, polypropylene, for extraction (0.45-jam filter), reservoir, and tared extraction
tubes
Rubber tubing, 3.2 IDx 1.6 ODx6.4 mm (1/8 IDx1/16ODx1 in) for connecting syringe
barrels.
Kjeltec Auto 2300 Sampler System, Tecator, Perstorp Analytical
Digestion tubes, straight neck, 250 ml_
Syringe filters, 0.45 urn, Whatman
Wash bottles
Vials, plastic
Centrifuge, Centra, GP-8, Thermo I EC, Needham Heights, MA
5.14.5 Reagents
Reverse osmosis deionized (RODI) water, ASTM Type I grade of reagent water
Ammonium acetate solution (NH4OAc), 1 N, pH 7.0. Add 1026 ml_ of glacial acetic acid
(CH3COOH) to 15 L RODI water. Add 1224 ml_ of concentrated ammonium hydroxide
(NH4OH). Cool. Allow to stand one day to equilibrate to room temperature. Mix and
adjust to pH 7.0 with CH3COOH (typically, ~ 40 ml_) or NH4OH and dilute with RODI water
to18L
Ethanol (CH3CH2OH), 95%, U.S.P.
Nessler's reagent. Add 4.56 g of potassium iodide (Kl) to 30 ml_ RODI water. Add 5.68 g
of mercuric iodide (Hgl2). Stir until dissolved. Dissolve 10 g of sodium hydroxide (NaOH)
in 200 mL of RODI water. Transfer NaOH solution to a 250-mL volumetric flask and slowly
add K-Hg-l solution. Dilute to volume with RODI water and thoroughly mix. Solution
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should not contain a precipitate. Solution can be used immediately. Store in brown bottle
to protect from light.
• Potassium chloride solution, 2 M. Add 1341.9 g of KCI reagent in 8 L RODI water. Allow
solution to equilibrate to room temperature. Dilute to 9 L with RODI water.
• Boric acid, 4% (w:v), with bromcresol green-methyl red indicator (0.075 % bromcresol
green and 0.05% methyl red), Chempure.
Hydrochloric acid (HCI), 0.1 N, standardized. Dilute 167 ml_ of concentrated HCI in 20 L of
RODI water. Refer to procedure for Standardization of Acids.
NaOH, 1 M. Add 500 ml_ of 50% NaOH solution to 8 L of RODI water. Dilute to 9 L with
RODI water.
5.14.6 Procedure
Extraction of Bases
1. Weigh 2.5 g of <2-mm, air-dry soil to the nearest mg and place in a labeled extraction tube
(ET). If sample is fine-grind, weigh 1 g to the nearest mg. If sample is moist, weigh enough
soil to achieve ~ 2.5 or 1g, respectively, of air-dry soil. Prepare one quality control check
sample per 24 samples.
2. Place labeled ET on extractor and connect to corresponding tared extraction tube
(TETNH4OAc) with rubber tubing.
3. Use wash bottle to rinse inside of ET with NH4OAc. All soil should be wetted and be sure
there are no air bubbles. Shaking, swirling, or stirring may be required to wet organic
samples. Fill ET to the 20-mL mark with NH4OAc solution (~ 10 ml_).
4. Secure reservoir tube (RT) to top of ET tube and let stand for 30 min. Extract at 30-min rate
the NH4OAc solution until 2 ml_ of this solution remains above soil level. Turn off extractor.
Do not let soil dry.
5. Add 40 mL of NH4OAc solution to the RT. Set extractor for an overnight (12h) extraction.
Extractor will turn off automatically.
6. Next day, remove RT from top of extractor and place in a clean container. Carefully remove
TETNH4OAc. Leave the rubber tubing on the ET. Weigh each TETNH4OAc containing the
NH4OAc extract to the nearest mg.
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7. Mix the extract in each TETNH4OAc by manually shaking. Fill a labeled plastic vial with
extract solution and cap. Discard the excess properly. The solution in the vial is reserved for
analyses of extracted cations (procedure 4B1a1b1-4) on the atomic absorption
spectrophotometer (AAS). Some samples may be cloudy and need to be filtered prior to
analysis on the AA. If extracts are not to be determined immediately after collection, then
store samples at 4°C in plastic tubes.
Removal of Excess Ammonium Acetate
8. Re-connect the TETNH4OAc with paired ET. Use a wash bottle to rinse the sides of the ET
with ethanol to remove any remaining NH4OAc or soil particles adhering to the ET. All soil
should be wetted and no air bubbles. Fill ET to the 20-mL mark with ethanol. Secure RT to
top of ET tube and let stand for 30 min.
9. Extract at the 30-min extraction rate the ethanol solution until 2 ml_ of this solution remains
above the soil level. Turn off the extractor. Do not let soil dry.
10. Add 45 ml_ of ethanol to the RT. Extract (« 45 min) the ethanol until 2 ml_ of this solution
remains above the soil level. Turn off the extractor. Do not let soil dry. Disconnect the
TETNH4OAc from the ET and discard the ethanol properly.
11. Re-connect the TETNH4OAc to the ET and add 55 mL of ethanol to the RT. Set the
extractor for 45 min. Turn off the extractor. Remove the TETNH4OAc, leaving the tubing
connected to the ET. Discard the ethanol properly.
12. After the final ethanol wash, collect a few drops of ethanol extract from the ET on a spot
plate. Test for NH4+ by using Nessler's reagent. A yellow, red to reddish brown precipitate
is a positive test. If the test is positive, repeat the ethanol wash and retest with Nessler's
reagent. Repeat until a negative test is obtained.
2 M KCI Rinse
13. Connect a new labeled extraction tube (ETKCI) with rubber tubing to ET on extractor.
14. Use a wash bottle to rinse inside of ET with 2 M KCI to remove any remaining ethanol or soil
particles adhering to the ET. All soil should be wetted and no air bubbles. Fill ET to the 20-
mL mark with KCI solution and let stand for 30.
15. Extract at the 30-min rate the KCI solution until 2 mL of this solution remains above soil
level. Turn off extractor. Do not let soil dry.
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16. Secure RT to top of ET tube. Add 40 ml_ KCI solution to RT and set the extract for 45 min.
Remove the ET and ETKCI from the extractor.
Steam Distillation: Setup, Operation, and Analysis
17. Transfer the contents of the ETKCI to a 250-mL digestion tube. If extracts are not to be
determined immediately after collection, then store samples at 4°C.
18. Refer to the manufacturer's manual for operation of the distillation unit. The following are
only very general guidelines for instrument conditions.
Program: Kjeldahl 1
Receiving solution (boric acid): 30 ml_
Water: 0 ml_
Alkali (NaOH): 20 ml_
Mode: Delay
Time: 1 s
Distillation: Volume
Tube Drain: Yes
19. When using new reagents, e.g., boric acid, reagent blanks are distilled in 2 sets of 6, one set
per Kjeltec machine. Each set of 6 is averaged and recorded on bench worksheet and
manually set on each machine. During the steam distillation, the mean reagent blank titer is
automatically subtracted from the sample titer.
20. Record the normality of standardized acid.
21. Connect the tube to the distillation unit. Close the safety door. Distillation and titration are
performed automatically. Record the titer in ml_ of titrant.
5.14.7 Calculations
CEC = [Titer x N x 100 x R]/[Sample Weight (g)]
where:
CEC = Cation Exchange Capacity (meq 100 g"1)
Titer = Titer of sample (ml_)
N = Normality of HCI titrant
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100 = Conversion factor to 100-g basis
R = Air-dry/oven-dry ratio (procedure 3D1) or field-moist/oven-dry ratio
(procedure 3D2)
5.14.8 Report
Report CEC-7 to the nearest 0.1 meq 100 g"1 (cmol (+) kg"1).
5.14.9 References
Holmgren, G.G.S., R.L. Juve, and R.C. Geschwender. 1977. A mechanically controlled variable
rate leaching device. Soil Sci. Am. J. 41:1207-1208.
Peech, M., L.T. Alexander, L.A. Dean, and J.F. Reed. 1947. Methods of soil analysis for soil
fertility investigations. USDA Circ. 757, 25 pp.
Sumner, M.E., and W.P. Miller. 1996. Cation exchange capacity and exchange coefficients, p.
1201-1229. In D.L Sparks (ed.) Methods of soil analysis. Part 3. Chemical methods. No.
5. ASA and SSSA, Madison, Wl.
5.15 Cation Exchange Capacity (Ca, Mg, K, Na)
5.15.1 Summary of Method
An NH4OAc extract (from section 5.13) is diluted with an ionization suppressant (La2O3). The
analytes are measured by an atomic absorption spectrophotometer (AAS). The analyte is
measured by absorption of the light from a hollow cathode lamp. An automatic sample changer
is used to aspirate a series of samples. The AAS converts absorption to analyte concentration.
Data are automatically recorded by a microcomputer and printer. The NH4OAc extracted
cations, Ca2+, Mg2+, K+, and Na+, are reported in meq 100 g"1 soil or (cmol (+) kg"1).
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5.15.2 Interferences
There are four types of interferences (matrix, spectral, chemical, and ionization) in the analyses
of these cations. These interferences vary in importance, depending upon the particular analyte
selected. Do not use borosilicate tubes because of potential leaching of analytes.
5.15.3 Precautions
Wear protective clothing and safety glasses. When preparing reagents, exercise special care.
Restrict the use of concentrated HCI to a fume hood. Many metal salts are extremely toxic and
may be fatal if ingested. Thoroughly wash hands after handling these metal salts.
Follow standard laboratory procedures when handling compressed gases. Gas cylinders
should be chained or bolted in an upright position. Acetylene gas is highly flammable. Avoid
open flames and sparks. Standard laboratory equipment includes fire blankets and
extinguishers for use when necessary. Follow the manufacturer's safety precautions when
using the AAS.
5.15.4 Equipment/Materials
• Electronic balance, ±1.0-mg sensitivity
• Atomic absorption spectrophotometer (AAS), double-beam optical system, AAnalyst, 300,
Perkin-ElmerCorp., Norwalk, CT
• Autosampler, AS-90, Perkin-Elmer Corp., Norwalk, CT
• Computer, with AA WinLab software, Perkin-Elmer Corp., Norwalk, CT, and printer
• Single-stage regulator, acetylene
• Digital diluter/dispenser, with syringes 10000 and 1000 uL, gas tight, MicroLab 500,
Hamilton Co., Reno, NV
• Plastic test tubes, 15-mL, 16 mm x 100, for sample dilution and sample changer
• Containers, polyethylene
• Peristaltic pump
5.15.5 Reagents
• Reverse osmosis deionized (RODI) water, ASTM Type I grade of reagent water
• Hydrochloric acid (HCI), concentrated 12 N
HCI, 1:1 HCI:RODI, 6 N. Carefully mix 1 part of concentrated HCI to 1 part RODI water.
• NH4OH, reagent-grade, specific gravity 0.90
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• Glacial acetic acid, 99.5%
• Ammonium acetate solution (NH4OAc), 1 N, pH 7.0. Add 1026 ml_ of glacial acetic acid
(CH3COOH) to 15 L RODI water. Add 1224 ml_ of concentrated ammonium hydroxide
(NH4OH). Cool. Allow to stand one day to equilibrate to room temperature. Mix and
adjust to pH 7.0 with CH3COOH (typically, « 40 ml_) or NH4OH and dilute with RODI water
to 18 L. The NH4OAc solution is used for extraction of cations (procedure 4B1a1).
NH4OAc solution, 2.0 N, pH 7.0. Mix 228 ml_ of glacial acetic acid in 1200 ml_ of RODI
water. While stirring, carefully add 272 ml_ of concentrated NH4OH. Cool. Allow to stand
one day to equilibrate to room temperature. Mix and adjust pH 7.0 using CH3COOH or
NH4OH. Dilute to 2 L with RODI water.
• Stock lanthanum ionization suppressant solution (SLISS), 65,000 mg L-1. Wet 152.4 g
lanthanum oxide (La2O3) with 100 mL RODI water. Slowly and cautiously add 500 ml_ of 6
N HCI to dissolve the La2O3. Cooling the solution is necessary. Dilute to 2 L with RODI
water. Filter solution. Store in polyethylene container.
• Working lanthanum ionization suppressant solution (WLISS), 2000 mg L-1. Dilute 61.5 mL
of SLISS with 1800 mL of RODI water (1:10). Dilute to final volume of 2-L with RODI
water. Store in polyethylene container.
• Primary stock standards solutions (PSSS), high purity, 1000 mg L-1: Ca, Mg, K, and Na.
• Working stock mixed standards solution (WSMSS), High, Medium, Low, Very Low, and
Blank. In five 500-mL volumetric flasks, add 250 mL of 2 N NH4 OAc and the following
designated amounts of Ca PSSS, Mg PSSS, K PSSS, and Na PSSS. Dilute to volume
with RODI. Invert to thoroughly mix. Store in polyethylene containers. Prepare fresh
weekly. Store in the refrigerator. Allow to equilibrate to room temperature before use.
Prepare WSMSS as follows:
o High Standard WSMSS: 90 mL Ca PSSS, 7.5 mL Mg PSSS, 20.0 mL K PSSS,
and 100.0 mL Na PSSS = 180 mg L"1 Ca, 15 mg L"1 Mg, 40 mg L"1 K, and 200 mg
L1 Na.
o Medium Standard WSMSS: 60 mL Ca PSSS, 5.0 mL Mg PSSS, 10.0 mL K
PSSS, and 50.0 mL Na PSSS = 120 mg L1 Ca, 10 mg L1 Mg, 20 mg L1 K, and
lOOmgL'1 Na.
o Low Standard WSMSS: 30 mL Ca PSSS, 2.5 mL Mg PSSS, 5.0 mL K PSSS,
and 10.0 mL Na PSSS = 60 mg L"1 Ca, 5 mg L"1 Mg, 10 mg L"1 K, and 20 mg L"1
Na.
o Low/Low Standard WSMSS: 12.5 mL Ca PSSS, 0.25 mL Mg PSSS, 0.125 mL K
PSSS, and 5.0 mL Na PSSS = 25 mg L1 Ca, 0.5 mg L1 Mg, 0.25 mg L1 K, and
lOmgL'1 Na.
o Blank WSMSS = 0 mL of Ca, Mg, K, and Na PSSS.
• Mixed calibration standard solutions (MCSS), High, Medium, Low, Very Low, and Blank.
Dilute 1 part WSMSS with 19 parts of WLISS (1:20) dilution with resulting concentrations
for MCSS as follows:
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o MCSS High Standard: 9.0 mg L"1 Ca, 0.75 mg L"1 Mg, 2.0 mg L"1 K, and 10.0 mg
L'1 Na.
o MCSS Medium Standard: 6.0 mg L"1 Ca, 0.5 mg L"1 Mg, 1.0 mg L"1 K, and 5.0
mg L"1 Na.
o MCSS Low Standard: 3.0 mg L"1 Ca, 0.25 mg L"1 Mg, 0.5 mg L"1 K, and 1.0 mg L"
1 Na.
o MCSS Very Low Standard: 1.25 mg L'1 Ca, 0.025 mg L'1 Mg, 0.125 K, and 0.5
mg L"1 Na.
o Blank = 0 mg L"1 Ca, Mg, K, and Na.
• Compressed air with water and oil traps.
• Acetylene gas, purity 99.6%.
5.15.6 Procedure
Dilution of Calibration Standards and Sample Extracts
1. The 10-mL syringe is for dilutent (WLISS). The 1-mL syringe is for the MCSS and NH4OAc
extracts (procedure 5.13). Set the digital diluter at a 1:20 dilution. See reagents for
preparation of the MCSS (High, Medium, Low, Very Low, Blank). Dilute 1 part NH4OAc
sample extract with 19 parts of WLISS (1:20 dilution).
2. Dispense the diluted sample solutions into test tubes which have been placed in the sample
holders of the sample changer.
AAS Set-up and Operation
Refer to the manufacturer's manual for operation of the AAS. The following are only very
general guidelines for instrument conditions for the various analytes.
Analyte
Ca
Mg
K
Na
Cone.
(mg L'1)
9.0
0.75
2.0
10.0
Burner &
angle
10cm @ 0°
10cm@0°
10cm @ 0°
10cm @ 30°
Wavelength
(nm)
422.7
285.2
766.5
589.0
Slit
(mm)
0.7
0.7
0.7
0.2
Fuel/Oxidant
(C2H2/Air)
1.5/10.0
1.5/10.0
1.5/10.0
1.5/10.0
3. Use the computer and printer to set instrument parameters and to collect and record
instrument readings.
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AAS Calibration and Analysis
4. Calibrate the instrument by using the MCSS (High, Medium, Low, Very Low, Blank). The
data system will then associate the concentrations with the instrument responses for each
MCSS. Rejection criteria for MCSS, if R2 <0.99.
5. If sample exceeds calibration standard, the sample is diluted 1:5, 1:20, 1:100, etc., with 1 N
NH4OAc followed by 1:20 dilution with WLISS.
6. Perform one quality control (QC) (Low Standard MCSS) every 12 samples. If reading is not
within 10%, the instrument is re-calibrated and QC re-analyzed.
7. Record analyte readings to 0.01 unit.
5.15.7 Calculation
The instrument readings for analyte concentration are in mg L"1. These analyte concentrations
are converted to meq 100 g"1 as follows:
Soil Analyte Concentration (meq 100 g"1) = [A x [(B1 - B2)/B3] x C x R x 100]/[1000 x E x F]
where:
A = Analyte (Ca, Mg, K, Na) concentration in extract (mg L"1)
B! = Weight of extraction syringe and extract (g)
B2 = Weight of tared extraction syringe (g)
B3 = Density of 1 N NH4OAc at 20°C (1.0124 g cm-3)
C = Dilution, if performed
100 = Conversion factor (100-g basis)
R = Air-dry/oven-dry ratio (procedure 3D1) or field-moist/oven-dry ratio
(procedure 3D2)
1000 = mLL-1
E = Soil sample weight (g)
F = Equivalent weight (mg meq"1)
where:
Ca+2 = 20.04 mg meq"1
Mg+2= 12.15 mg meq"1
Na+1 = 22.99 mg meq'1
K+1 = 39.10mgmeq-1
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5.15.8 Reporting
Report the extractable Ca2+, Mg2+, Na+, and K+ to the nearest 0.1 meq 100 g"1 (cmol (+) kg"1).
5.15.9 References
Thomas, G.W. 1982. Exchangeable cations, p. 159-165. In A.L Page, R.H. Miller, and D.R.
Keeney (eds.) Methods of soil analysis. Part 2. Chemical and microbiological properties.
2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, Wl.
5.16 Acid Oxalate Extraction (Al, Fe, Mn, P, Si)
5.16.1 Summary of Method
A soil sample is extracted with a mechanical vacuum extractor (Holmgren et al., 1977) in a 0.2
M ammonium oxalate solution buffered at pH 3.0 under darkness. The ammonium oxalate
extract is weighed. The ammonium oxalate extract is diluted with reverse osmosis deionized
water. The analytes are measured by a inductively coupled plasma atomic emission
spectrophotometer (ICP-AES). Data are automatically recorded by a computer and printer. All
these data are reported in percent except Mn and P, which are reported in mg kg"1.
5.16.2 Interferences
There are four types of interferences (matrix, spectral, chemical, and ionization) in the ICP
analyses of these elements. These interferences vary in importance, depending upon the
particular analyte chosen.
The ammonium oxalate buffer extraction is sensitive to light, especially UV light. The exclusion
of light reduces the dissolution effect of crystalline oxides and clay minerals. If the sample
contains large amounts of amorphous material (>2% Al), an alternate method should be used,
i.e., shaking with 0.275 M ammonium oxalate, pH 3.25, 1:100 soil:extractant.
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5.16.3 Precautions
Wear protective clothing and eye protection. When preparing reagents, exercise special care.
Follow standard laboratory practices when handling compressed gases. Gas cylinders should
be chained or bolted in an upright position. Follow the manufacturer's safety precautions when
using the UV spectrophotometer and ICP.
5.16.4 Equipment/Materials
• Electronic balance, ±1.0-mg sensitivity
• Mechanical vacuum extractor, 24 place, SAMPLETEX, MAVCO Industries, Lincoln, NE
• Tubes, 60-mL, polypropylene, for extraction (0.45-um filter), reservoir, and tared extraction
tubes
• Rubber tubing, 3.2 IDx 1.6ODx6.4 mm, (1/8 IDx 1/16 OD x 1 in) for connecting syringe
barrels
• Dispenser, 30-mL
• Pipettes, electronic digital, 10000 and 1000 uL, with tips 10000 and 1000 uL
• Containers, polyethylene
• Inductively coupled plasma atomic emission spectrophotometer (ICP-AES), dual-view, with
high-solids nebulizer, alumina or quartz injector, Optima 4300 DV, Perkin-Elmer Corp.,
Norwalk, CT.
• Automsampler, AS-93, Perkin-Elmer Corp., Norwalk, CT
• Computer, with Winl_ab32TM software, and printer
• Single-stage regulator, high-purity, high-flow, argon
• Test tubes, 15-mL, 16 mm x 100, for sample dilution and sample changer, Fisher Scientific
• Vortexer, mini, MV1, VWR Scientific Products
• Spectrophotometer, UV-Visible, Varian, Gary 50 Cone, Varian Australia Pty Ltd.
• Computer with Gary WinUV software, Varian Australia Pty Ltd., and printer
• Cuvettes, plastic, 4.5-mL, 1-cm light path, Daigger Scientific
5.16.5 Reagents
• Reverse osmosis deionized (RODI) water, ASTM Type I grade of reagent water
• Ammonium oxalate buffer solution, 0.2 M, pH 3.0. Solution A (base): Dissolve 284 g of
(NH4)2C2Cv2H2O in 10 L of Dl water. Solution B (acid): Dissolve 252 g of h^CU-hbO in
10 L of Dl water. Mix 4 parts solution A with 3 parts solution B. Adjust ammonium oxalate
solution pH by adding either acid or base solution. Store in a polypropylene bottle.
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• Borax pH buffers, pH 4.00, 7.00, and 9.18 for electrode calibration, Beckman, Fullerton,
CA.
• Primary Fe standard, 1000 mg L-1. Certified Reference Solution, Fisher Chemical
Scientific Co., Fairlawn, N.J.
• Primary Al standard, 1000 mg L-1. Certified Reference Solution, Fisher Chemical Scientific
Co., Fairlawn, N.J.
• Primary Si standard, 1000 mg L-1. Certified Reference Solution, Fisher Chemical Scientific
Co., Fairlawn, N.J.
• Primary Mn standard, 1000 mg L-1. Certified Reference Solution, Fisher Chemical
Scientific Co., Fairlawn, N.J.
• Primary P standard, 1000 mg L-1. Certified Reference Solution, Fisher Chemical Scientific
Co., Fairlawn, N.J.
• High mixed calibration standard. Mix 60 mL of each primary standard (Si, Fe, and Al) with
10 mL of primary Mn standard and 20 mL of primary P standard in 1 L volumetric flask.
Add 100 mL of 0.2 M ammonium oxalate solution and make to 1-L volume with RODI
water. The elements are added in the order (Si, Fe, Al, Mn, and P) to avoid element
precipitation. Resulting solution contains 60 mg L-1 each of Si, Fe, and Al, 10 mg L-1 Mn,
and 20 mg L-1 P. Invert to mix thoroughly. Store in a polyethylene bottle. Make fresh
weekly. Store in a refrigerator.
• Medium mixed calibration standard. Mix 30 mL of each primary standard (Si, Fe, and Al)
with 5 mL of primary Mn standard and 10 mL of primary P standard in 1 L volumetric flask.
Add 100 mL of 0.2 M ammonium oxalate solution and make to 1-L volume with RODI
water. The elements are added in the order (Si, Fe, Al, Mn, and P) to avoid element
precipitation. Resulting solution contains 30 mg L-1 each of Si, Fe, and Al, 5 mg L-1 Mn,
and 10 mg L-1 P. Invert to mix thoroughly. Store in a polyethylene bottle. Make fresh
weekly. Store in a refrigerator.
• Low mixed calibration standard. Mix 10 mL of each primary standard (Si, Fe, and Al) with
2 mL of primary Mn standard, and 3 mL primary P standard in 1 L volumetric flask. Add
100 mL of 0.2 M ammonium oxalate solution and make to 1-L volume with RODI water.
The elements are added in the order (Si, Fe, Al, Mn, and P) to avoid element precipitation.
Resulting solution contains 10 mg L-1 each of Si, Fe, and Al, 2 mg L-1 Mn, and 3 mg L-1
P. Invert to mix thoroughly. Store in a polyethylene bottle. Make fresh weekly. Store in a
refrigerator.
• Low Si calibration standard. Mix 5 mL of Si primary standard in 1-L volumetric flask. Add
100 mL of 0.2 M ammonium oxalate solution and make to 1-L volume with RODI water.
Resulting solution contains 5 mg L-1 Si. Invert to mix thoroughly. Store in polyethylene
bottle. Make fresh weekly. Store in a refrigerator.
• Very Low Si calibration standard. Mix 2 mL of Si primary standard in 1-L volumetric flask.
Add 100 mL of 0.2 M ammonium oxalate solution and make to 1-L volume with RODI
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water. Resulting solution contains 2 mg L-1 Si. Invert to mix thoroughly. Store in
polyethylene bottle. Make fresh weekly. Store in a refrigerator.
• Calibration reagent blank solution. Add 100 ml_ of 0.2 M ammonium oxalate solution and
make to 1-L volume with RODI water. Store in polyethylene bottle. Make fresh weekly.
Store in a refrigerator.
• Argon gas, purity 99.9%
• Nitrogen, purity 99.9%
5.16.6 Procedure
Extraction of Fe, Mn, Al, Si, and P
Weigh 0.5 g of <2-mm, air-dry or fine-grind soil the nearest mg and place in sample tube. If
sample is moist, weigh enough soil to achieve ~ 0.5 g. Prepare two reagent blanks (no sample
in tube) per set of 48 samples.
1. Place labeled ET on extractor and connect to corresponding tared extraction tube
(TETOxalate) with rubber tubing.
2. Use a dispenser to add 15.00 ml_ of ammonium oxalate buffer to the ET. Make sure that the
sample is thoroughly wetted. During the addition, wash sides of the tube and wet the
sample. Shaking, swirling, or stirring may be required to wet organic samples. Allow
sample to stand for at least 30 min. Cover samples with black plastic bag to exclude light.
3. Secure reservoir tube (RT) to top of ET tube. Set extractor for 30-min extraction rate and
extract until the ammonium oxalate buffer solution is at a 0.5 to 1.0-cm height above
sample. Turn off extractor.
4. Add 35 mL of ammonium oxalate buffer to the RT.
5. Cover the extractor with a black plastic bag to exclude light. Adjust the extraction rate for a
12-h extraction.
6. After the extraction, shut off the extractor. Carefully remove TETOxalate. Leave the rubber
tubing on the ET.
7. Weigh each syringe containing ammonium oxalate extract to the nearest mg.
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8. Mix extract in each TETOxalate by manually shaking. Fill a disposable tube with extract
solution. This solution is reserved for determinations of Fe, Mn, Al, Si, and P. If optical
density is to be measured, fill a disposable cuvette with extract solution. Discard excess
solution properly. If extracts are not to be determined immediately after collection, then
store samples at 4°C.
Determination of Optical Density of Extract
9. Place 4 ml_ of ammonium oxalate extract in disposable cuvette.
10. Place 4 ml_ of ammonium oxalate reagent blank in disposable cuvette.
11. On the spectrophotometer, select a 430-nm wavelength. Select normal slit width and
height. Refer to manufacturer's manual for operation of the spectrophotometer.
12. Use the ammonium oxalate extract reagent blank to zero spectrophotometer.
13. Record optical density of ammonium oxalate extract to nearest 0.000 unit.
Dilution of Sample Extracts and Standards
14. Dilute ammonium oxalate extracts (1:10) with RODI water. Add 1 part ammonium oxalate
sample extract with 9 parts dilution solution. Pipet 0.7 ml_ of extract and 6.3 ml_ RODI
water. Vortex. Calibration reagent blanks and calibration standards are not diluted.
15. Dispense the diluted solutions into test tubes that have been placed in the sample holder of
the sample changer.
ICP-AES Set-up and Operation
16. Refer to the manufacturer's manual for operation of the ICP-AES. The following
parameters are only very general guidelines for instrument conditions for the various
analytes.
Parameter Value
Plasma
Source Equilibration Delay 20 sec
Plasma Aerosol Type Wet
Nebulizer Start-up Conditions Gradual
Plasma 15Lmin"1
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Auxiliary
Nebulizer
Power
View Dist
Plasma View
Peristaltic Pump
Sample Flow Rate
Sample Flush Time
Wash Parameters
Wash Rate
Wash Time
Background Correction
Read Delay
Replicates
0.5 Lmin"1
0.85 L min"1
1450 Watts
15.0
Radial
2.00 L mirT1
35 sec
Between samples
2.00 mL min"1
30 sec
2 point (all elements)
2 sec
2
Nebulizer pressure depends on the type of nebulizer that is being used, i.e., low flow
nebulizer requires a higher nebulizer pressure whereas a higher flow nebulizer requires a
lower nebulizer pressure. To check for correct nebulizer pressure, aspirate with 1000.0 mg
L"1 yttrium. Adjust pressure to correct yttrium bullet.
17. Analyte data are reported at the following wavelengths:
Element
Wavelength (nm)
Fe
Al
Si
Mn
P
259.94
308.22
251.61
257.61
213.61
18. Use the computer and printer to set instrument parameters and to collect and record
instrument readings. The instrument readings are programmed in mg L"1.
ICP-AES Calibration and Analysis
19. Use a multipoint calibration for ICP-AES analysis of ammonium oxalate extracts. The ICP
calibrates the blank first, low standard, medium standard, followed by the high standard.
Prepare a quality control (QC) standard with analyte concentration between the high and
low calibration standards. The ICP reads the QC after the high standard. If the QC falls
within the range set by operator (±10%), the instrument proceeds to analyze the unknowns.
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If the QC is outside the range, the instrument restandardizes. The QC is analyzed
approximately every 12 samples.
20. If sample exceeds calibration standard, dilute 1:5 (1 ml_ of sample extract with 4 ml_ 0.02 M
ammonium oxalate extracting solution), followed by a 1:10 (1 mL of 1:5 solution with 9 mL
RODI water). This makes for a 1:50 dilution.
21. Record analyte readings to the nearest 0.01 unit.
5.16.7 Calculation
The instrument readings are the analyte concentration (mg L"1 Fe, Mn, Al, Si, and P). Use these
values to calculate the analyte concentration in percent in the soil for Fe, Al, and Si and mg kg"1
for Mn and P as follows:
Soil Fe, Al, Si (%) = [A x [(B1 - B2)/B3] x C1 x C2 x R x 100]/(E x 1000 x 1000)
where:
A = Sample extract reading (mg L"1)
B! = Weight of syringe + extract (g)
B2 = Tare weight of syringe (g)
B3 = Density of 0.2 M ammonium oxalate solution at 20° C (1.007 g mL"1)
d = Dilution, required
C2 = Dilution, if performed
R = Air-dry/oven-dry ratio or field-moist/oven-dry ratio
E = Sample weight (g)
100 = Conversion factor to 100-g basis
1000 = Factor in denominator (mL L"1)
1000 = Factor in denominator (mg g"1)
Soil Mn, P (mg kg"1) = [ A x [(B-^ - B2)/B3] x d x C2 x R x 1000]/(E x 1000)
where:
A = Sample extract reading (mg L"1)
B! = Weight of syringe + extract (g)
B2 = Tare weight of syringe (g)
B3 = Density of 0.2 M ammonium oxalate solution at 20° C (1.007 g mL"1)
d = Dilution, required
C2 = Dilution, if performed
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R = Air-dry/oven-dry ratio or field-moist/oven-dry ratio
1000 = Conversion factor in numerator to kg-basis
1000 = Factor in denominator (ml_ L"1)
E = Sample weight (g)
5.16.8 Reporting
Report the percent ammonium oxalate extractable Al, Fe, and Si to the nearest 0.01%. Report
the concentration of ammonium oxalate extractable Mn and P to the nearest mg kg"1 soil.
Report the optical density of the ammonium oxalate extract to the nearest 0.01 unit.
5.16.9 References
Aguilera, N.H., and M.L. Jackson. 1953. Iron oxide removal from soils and clays. Soil Sci. Soc.
Am. Proc. 17:359-364.
Holmgren, G.G.S., R.L. Juve, and R.C. Geschwender. 1977. A mechanically controlled variable
rate leaching device. Soil Sci. Am. J. 41:1207-1208.
Jackson, M.L. 1979. Soil chemical analysis-advance course. 2nd ed., 11th Printing. Published
by author, Madison, Wl.
Mehra, O.P., and M.L. Jackson. 1960. Iron oxide removal from soils and clays by a dithionite-
citrate system buffered with sodium bicarbonate. Clays Clay Miner. 7:317-327.
Soil Survey Staff. 1999. Soil taxonomy. A basic system of soil classification for making and
interpreting soil surveys. 2nd ed. USDA-NRCS. Govt. Print. Office, Washington DC.
Wada, K. 1989. Allophane and imogolite. p. 1051-1087. In J.B. Dixon and S.B. Weed (eds.).
Minerals in soil environment. 2nd ed. Soil Sci. Soc. Am. Book Series No. 1. ASA and
SSSA, Madison.
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5.17 Electrical Conductivity (Saline Prediction)
5.17.1 Summary of Method
A soil sample is mixed with water and allowed to stand overnight. The electrical conductivity
(EC) of the mixture is measured using an electronic bridge. The EC by this method (4F1a1a1)
is used to indicate the presence of soluble salts (U.S. Salinity Laboratory Staff, 1954).
5.17.2 Interferences
Reverse osmosis deionized water is used to zero and flush the conductivity cell. The extract
temperature is assumed to be 25°C. If the temperature deviates significantly, a correction may
be required.
Provide airtight storage of KCI solution and samples to prevent soil release of alkali-earth
cations. Exposure to air can cause gains and losses of water and dissolved gases significantly
affecting EC readings.
5.17.3 Precautions
No significant hazards are associated with this procedure. Follow standard laboratory safety
practices.
5.17.4 Equipment/Materials
• Electronic balance, ±1.0-mg sensitivity
• Conductivity bridge and conductivity cell, with automatic temperature adjustment, 25 ±
0.1°C, Markson Model 1056, Amber Science, Eugene, OR
• Plastic cups, 30 ml_ (1 fl. oz.), with lids, Sweetheart Cup Co. Inc., Owings Mills, MD
• Dispenser, Repipet or equivalent, 0 to 10 mL
5.17.5 Reagents
• Reverse osmosis (RO) water, ASTM Type III grade of reagent water
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• Potassium chloride (KCI), 0.010 N. Dry KCI overnight in oven (110°C). Dissolve 0.7456 g
of KCI in RODI water and bring to 1-L volume. Conductivity at 25°C is 1.412 mmhos cm"1.
5.17.6 Procedure
1. Weigh 5.0 g of <2-mm, air-dry soil in a 30-mL (1-oz) condiment cup.
2. Add 10 mL of RO water to sample using a Repipet dispenser.
3. Swirl to mix, cap, and allow to stand overnight.
4. Standardize the conductivity bridge using RO water (blank) and 0.010 N KCI (1.41 mmhos
cm'1).
5. Read conductance of supernatant solution directly from the bridge.
6. Record conductance to 0.01 mmhos cm"1.
5.17.7 Calculation
1. No calculations are required for this procedure.
2. Use the following relationship to estimate the total soluble cation or anion concentration
(meq L"1) in the soil.
3. EC (mmhos cm-1) x 10 = Cation or Anion (meq L"1)
4. Use the following relationship to estimate the total soluble cation or anion concentration
(meq g"1 oven-dry soil) in the soil.
EC (mmhos cm"1) x 20 = Cation (meq g"1 soil)
EC (mmhos cm"1) x 20 = Anion (meq g"1 soil)
5.17.8 Reporting
Report prediction conductance to the nearest 0.01 mmhos cm"1 (dS m"1).
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5.17.9 References
U.S. Salinity Laboratory Staff. 1954. L.A. Richards (ed.) Diagnosis and improvement of saline
and alkali soils. 160 p. USDA Handb. 60. U.S. Govt. Print. Office, Washington, DC.
5.18 Citrate Dithionite Extraction (Al, Fe, and Mn)
5.18.1 Summary of Method
A soil sample is mixed with sodium dithionite, sodium citrate, and reverse osmosis deionized
(RODI) water, and shaken overnight. Solution is centrifuged, and a clear extract obtained. The
CD extract is diluted with RODI water. The analytes are measured by an atomic absorption
spectrophotometer (AAS). The data are automatically recorded by a computer and printer. The
AAS converts absorption to analyte concentration.
5.18.2 Interferences
There are four types of interferences (matrix, spectral, chemical, and ionization) in the AA
analyses of these elements. These interferences vary in importance, depending upon the
particular analyte selected.
The redox potential of the extractant is dependent upon the pH of the extracting solution and the
soil system. Sodium citrate complexes the reduced Fe and usually buffers the system to a pH
of 6.5 to 7.3. Some soils may lower the pH, resulting in the precipitation of Fe sulfides.
Filtered extracts can yield different recoveries of Fe, Mn, and Al, relative to unfiltered extracts.
5.18.3 Precautions
Wear protective clothing (coats, aprons, sleeve guards, and gloves); eye protection (face
shields, goggles, or safety glasses); and a breathing filter when handling dry sodium dithionite.
Sodium dithionite may spontaneously ignite if allowed to become moist, even by atmospheric
moisture. Keep dithionite in a fume hood.
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Follow standard laboratory practices when handling compressed gases. Gas cylinders should
be chained or bolted in an upright position. Acetylene gas is highly flammable. Avoid open
flames and sparks. Standard laboratory equipment includes fire blankets and extinguishers for
use when necessary. Follow the manufacturer's safety precautions when using the AAS.
5.18.4 Equipment/Materials
• Electronic balance, ±1.0-mg sensitivity
• Mechanical reciprocating shaker, 200 oscillations min-1, 1 1/4 in strokes, Eberbach 6000,
Eberbach Corp., Ann Arbor, Ml
• Atomic absorption spectrophotometer (AAS), double-beam optical system, AAnalyst, 300,
Perkin-ElmerCorp., Norwalk, CT, with computer and printer
• Autosampler, AS-90, Perkin-Elmer Corp., Norwalk, CT
• Peristaltic pump
• Single-stage regulators, acetylene and nitrous oxide
• Centrifuge, Centra, GP-8, Thermo IEC, Needham Heights, MA
• Digital diluter/dispenser, with syringes 10000 and 1000 uL, gas tight, MicroLab 500,
Hamilton Co., Reno, NV
• Dispenser, 30 ml_
• Test tubes, 15-mL, 16 mm x 100, for sample dilution and sample changer
• Containers, polypropylene
• Volumetrics, Class A, 100, 250, and 1000-mL
• Measuring scoop, handmade, 0.4 g calibrated
• Centrifuge tubes, 50-mL
5.18.5 Reagents
• Reverse osmosis deionized (RODI) water, ASTM Type I grade of reagent water
• Sodium dithionite (Na2S2O4), purified powder
• Sodium citrate dihydrate (Na3C6H507«2H2O), crystal, reagent. Dissolve 336 g sodium citrate
in approximately 1 L RODI water, followed by diluting to 2 L with RODI water. Final
concentration is 0.57 M sodium citrate.
• Sulfuric acid (H2SO4), concentrated.
• Phosphoric acid (H3PO4), concentrated (85%). For Fe analysis, samples are diluted 1:50
prior to analysis. A diluting solution for Fe analysis (for a final concentration of 0.5%
H3PO4 in samples) may be made by adding 6.12 ml_ of concentrated H3PO4 to 500 ml_
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volume of RODI water, diluting to 1000 ml_ and mixing thoroughly. (Note: 1:5 sample
dilutions for Al and Mn are in RODI water.)
• Primary Stock Standard Solution (PSSS), high purity, 1000 mg L-1: Fe, Mn, and Al.
• Calibration standards for Fe (See next bullet): To each 100 ml_ volume of blank,
calibration, and quality control (QC) standards, add 25 ml_ of the following matrix matching
mixture. This mixture is made by combining 20 ml_ Na citrate extracting solution, 0.21 mL
H2SO4, and 6 ml_ H3PO4 and diluting to 250 ml_ volume with RODI water. Invert to mix
thoroughly. (Note: Matrix of standards is prepared to match a 1:50 dilution of samples.
Also, H2SO4 substitutes for dithionite).
Standard Fe calibration solutions (SFeCS) or working standards (25.0, 20.0, 15.0, 10.0,
5.0, 1.0, and 0.0 mg Fe L-1) and QC (12.5 mg L-1). Prepare fresh weekly. In seven 100-
mL volumetric flasks add as follows:
o 25.0 mg Fe L'1 = 2.5 mL PSSSFe
o 20.0 mg Fe L'1 = 2.0 mL PSSSFe
o 15.0 mg FeL'1 = 1.5 mL PSSSFe
o 10.0 mg FeL'1 = 1.0 mL PSSSFe
o 5.0 mg Fe L"1 = 0.5 mL PSSSFe
o 1.0 mg Fe L'1 = 0.1 mL PSSSFe
o 0.0 mg Fe L'1 = 0.0 mL PSSSFe (blank)
o 12.5 m Fe L'1 = 1.25 mL PSSSFe (QC)
• Fill to volume with RODI water and invert to mix thoroughly. After dissolution, transfer
solution to a plastic bottle.
• Calibration standards for Mn (See next bullet): To each 100 mL volume of blank,
calibration, and quality control (QC) standards, add 25 mL of the following matrix matching
mixture. This mixture is made by combining 200 mL Na citrate extracting solution, 2.1 mL
H2SO4, and diluting to 250-mL volume with RODI water. Invert to mix thoroughly. (Note:
Matrix of standards is prepared to match a 1:5 dilution of samples. Also, H2SO4 substitutes
for dithionite).
• Standard Mn calibration solutions (SMnCS) or working standards (15.0, 10.0, 5.0, 2.5, 1.5,
and 0.0 mg Mn L-1) and QC (6.5 mg L-1). Prepare fresh weekly. In six 100-mL volumetric
flasks add as follows:
o 15.0 mgMn L'1 = 1.5mLPSSSMn
o 10.0 mgMn L'1 = 1.0mL PSSSMn
o 5.0 mg Mn L'1 = 0.5 mL PSSSMn
o 2.5 mg Mn L'1 = 0.25 mL PSSSMn
o 1.5 mgMnL'1 = 0.15 mL PSSSMn
o O.Omg Mn L"1 = 0.0 mL PSSSMn (blank)
o 6.5 mg Mn L1 = 0.65 mL PSSSMn (QC)
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• Fill to volume with RODI water and invert to mix thoroughly. After dissolution, transfer
solution to a plastic bottle.
• Calibration standards for Al (See next bullet): To each 100 ml_ volume of blank, calibration,
and quality control (QC) standards, add 25 ml_ of the following matrix matching mixture.
This mixture is made by combining 200 ml_ Na citrate extracting solution, 2.1 ml_ H2SO4,
and then diluting to 250-mL volume with RODI water. Invert to mix thoroughly. (Note:
Matrix of standards is prepared to match a 1:5 dilution of samples (same as with Mn).
Also, H2SO4 substitutes for dithionite).
• Standard Al calibration solutions (SAICS) or working standards (100.0, 80.0, 60.0, 40.0,
20.0, 10.0, and 0.0 mg Al L-1) and QC (50.0 mg L-1). Prepare fresh weekly. In seven 100-
ml_ volumetric flasks add as follows:
o 100.0 mg Al L'1 = 10.0 ml_ PSSSAI
o 80.0 mg Al L'1 = 8.0 ml_ PSSSAI
o 60.0 mg Al L'1 = 6.0 ml_ PSSSAI
o 40.0 mg Al L'1 = 4.0 ml_ PSSSAI
o 20.0 mg Al L'1 = 2.0 ml_ PSSSAI
o 10.0 mg Al L'1 = 1.0 ml_ PSSSAI
o 0.0 mg Al L'1 = 0.0 ml_ PSSSAI (blank)
o 50.0 mg Al L'1 = 5.0 ml_ PSSSAI (QC)
• Fill to volume with RODI water and invert to mix thoroughly. After dissolution, transfer
solution to a plastic bottle.
• Acetylene gas, purity 99.6%
• Nitrous oxide, USP
• Compressed air with water and oil traps
5.18.6 Procedure
Extraction of Al, Fe, and Mn
1. Weigh 0.75 g of <2-mm or fine-grind, air-dry soil sample to the nearest mg and place in an
50-mL centrifuge tube. If sample is moist, weigh enough soil to achieve « 0.75 g of air-dry
soil.
2. Add 0.4 g of sodium dithionite (use one calibrated scoop) and 25 ml_ of sodium citrate
solution.
3. Cap tubes and shake briefly by hand to dislodge soil from tube bottom. Place tubes in rack.
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4. Place rack in shaker and shake overnight (12 to 16 h) at 200 oscillations min"1 at room
temperature (20°C ± 2°C).
5. Remove tubes from shaker and manually shake tubes to dislodge any soil from cap. Allow
samples to sit overnight.
6. The following day, centrifuge at 4000 rpm for 15 min. The Fe, Mn, and Al are determined on
the AAS from a clear aliquot of solution.
Dilution of Sample Extracts
No ionization suppressant is required as the Na in the extractant is present in sufficient quantity.
For a 1:50 dilution of samples for Fe analysis, use the H3PO4 diluting solution (see Section
5.9.5, bullet #5). The dilution of Fe results in a final solution concentration of 0.5% H3PO4.
Dilute 1 part CD sample extract with 49 parts of H3PO4 diluting solution (1:50 dilution).
7. A 1:5 dilution in RODI water is used for Al and Mn. Dilute 1 part CD sample extract with 4
parts RODI water.
8. Dispense the diluted sample solutions into test tubes that have been placed in the holders of
the sample changer.
AAS Set-up and Operation
Refer to the manufacturer's manual for operation of the AAS. The following are only very
general guidelines for instrument conditions for the various analytes.
Analyte
Fe
Mn
Al
Cone.
(mg L-1)
25.0
15.0
100.0
Burner Head
10-cm parallel
10-cm parallel
5-cm parallel
Wavelength
(nm)
248.8
279.8
309.3
Slit
(mm)
0.2
0.2
0.7
Fuel/Oxidant
(C2H2/Air)
3.0/15.7
3.0/15.7
8.5/15.7
Typical read delay is 3 s, and integration time is 3 s but can vary depending on soil type. Three
replicates are average for each sample.
9. Use the computer and printer to set instrument parameters and to collect and record
instrument readings.
10. The instrument readings are programmed to display analyte concentration in mg L"1 (ppm).
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AAS Calibration and Analysis
Each element is analyzed during separate runs on the AAS. Use the calibration reagent blank
and calibration standards to calibrate the AAS. Calibrations are linear with calculated intercept.
11. Use the QC after every 12th sample. It must pass within 15% to continue. If it fails,
recalibrate and reread the QC. The QC is also read at the end of each run.
12. If samples are outside the calibration range, a serial dilution is performed. A 1:5 dilution of
the sample using the calibration blank, followed by the typical dilution (1:5 dilution with RODI
water for Al and Mn, and 1:50 dilution with the H3PO4 diluting solution for Fe). Maintain
matrix match between standards and diluted samples by performing this extra dilution with
calibration blank.
13. Record analyte readings to 0.01 unit.
5.18.7 Calculation
Convert analyte concentrations (mg L"1) to percent in soil as follows:
Soil Fe, Al, Mn (%) = (A x B x C x R x 100)/ (E x 1000)
where:
A = Sample extract reading (mg L~1)
B = Extract Volume (L)
C = Dilution, required
R = Air-dry/oven-dry ratio (procedure 3D1) or field-moist/oven-dry ratio
(procedure 3D2)
E = Sample weight (g)
100 = Conversion factor to 100-g basis
1000 = mg g"1
5.18.8 Reporting
Report percent CD extractable Al, Fe, and Mn to the nearest 0.1 of a percent.
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5.18.9 References
Wada, K. 1989. Allophane and imogolite. p. 1051-1087. In J.B. Dixon and S.B. Weed (eds.).
Minerals in soil environment. 2nd ed. Soil Sci. Am. Book Series No. 1. ASA and SSSA,
Madison, Wl.
Soil Survey Staff. 1999. Soil taxonomy: A basic system of soil classification for making and
interpreting soil surveys. USDA-NRCS Agric. Handb. 436. 2nd ed. U.S. Govt. Print.
Office, Washington, DC.
5.19 Phosphorus (Olsen)
5.19.1 Summary of Method
A 1.0-g soil sample is shaken with 20 ml_ of Olsen sodium-bicarbonate extracting solution for 30
min. The sample is centrifuged until solution is free of soil mineral particles, and then filtered
until clear extracts are obtained. Dilute 5-mL of sample extract with 5-mL of color reagent. The
absorbance of the solution is read using a spectrophotometer at 882 nm. Data are reported as
mg P kg"1 soil.
5.19.2 Interferences
The Mo blue methods, which are very sensitive for P, are based on the principle that in an acid
molybdate solution containing orthophosphate ions, a phosphomolybdate complex forms that
can be reduced by ascorbic acid, SnCI2, and other reducing agents to a Mo color. The intensity
of blue color varies with the P concentration but is also affected by other factors such as acidity,
arsenates, silicates, and substances that influence the oxidation-reduction conditions of the
system (Olsen and Sommers, 1982).
5.19.3 Precautions
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses). When preparing reagents, exercise special care. Many
metal salts are extremely toxic and may be fatal if ingested. Thoroughly wash hands after
handling these metal salts. Restrict the use of concentrated H2SO4 and HCI to a fume hood.
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Use safety showers and eyewash stations to dilute spilled acids and bases. Use sodium
bicarbonate and water to neutralize and dilute spilled acids.
5.19.4 Equipment/Materials
Electronic balance, ±1.0-mg sensitivity
Mechanical reciprocating shaker, 200 oscillations min-1, 1 1/2 in strokes, Eberbach 6000,
Eberbach Corp., Ann Arbor, Ml
Centrifuge tube, 50 ml_, polyethylene
Funnel, 60° angle, long stem, 50-mm diameter
Filter paper, Whatman 42, 150 mm
Centrifuge, Centra, GP-8, Thermo IEC, Thermo IEC, Needham Heights, MA.
Pipettes, electronic digital, 1000uL and 10 ml_, with tips, 1000uL and 10 ml cups, plastic
Dispenser, 30 ml_ or 10 mL
Cuvettes, plastic, 4.5-mL, 1-cm light path, Daigger Scientific
Spectrophotometer, UV-Visible, Dual-View, Varian, Gary 50 Cone, Varian Australia Pty Ltd.
Computer, with Gary WinUV software, Varian Australia Pty Ltd., and printer
5.19.5 Reagents
Sulfuric acid (H2SO4), concentrated, 36 N, trace pure grade
Sulfuric acid (H2SO4), 4 M. To 250-mL volumetric, carefully add 56 mL concentrated H2SO4
to 150 mL RODI water. Allow to cool. Make to final volume with RODI water. Invert to
thoroughly mix.
NaOH, 1 M. Dissolve 4 g NaOH in 100 mL RODI water.
Olsen Sodium Bicarbonate Extracting solution (0.5 M NaHCO3). To 6-L container, dissolve
252 g NaHCO3 in RODI water. Adjust the pH to 8.5 with 1M NaOH. Make to final volume.
Mix thoroughly. Check pH every day.
Ammonium molybdate, 4%. Dissolve 4 g of [(NH4)6Mo7O24'4H2O] in 100-mL volumetric
with RODI water. Dilute to volume with RODI water. Store in the dark in the refrigerator.
Potassium antimony- (III) oxide tartarate, 0.275%. Dissolve 0.275 g
[K(SbO)C4H4O6'1/2H2O] in 100 mL RODI water.
Ascorbic acid, 1.75%. Dissolve 1.75 g ascorbic acid in 100 mL RODI water. Prepare fresh
daily.
Color developing reagent. To a 500-mL bottle, add 50 mL 4 M H2SO4, 15 mL 4%
ammonium molybdate, 30 mL 1.75% ascorbic acid, 5 mL 0.275% potassium antimony-
(III) oxide tartarate, and 200 mL RODI water. Mix well after each addition. Prepare fresh
daily.
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• Stock standard P solution (SSPS), 100.0 mg P L-1. In a 1-L volumetric flask, dissolve
0.4394 g primary standard grade anhydrous potassium dihydrogen phosphate (KH2PO4)
that has been dried for 2 h at 110°C in about 800 ml_ extracting solution. Dilute to 1-L
volume with extracting solution and invert to thoroughly mix. Store in polyethylene
containers. Make fresh weekly. Store in a refrigerator.
Working stock standard P solution (WSSPS), 4.0 mg P L-1. Pipet 10 mL of 100 mg P L-1
SSPS to 250-mL volumetric flask. Dilute to 250-mL volume with extracting solution and
invert to thoroughly mix. Make fresh weekly. Store in the refrigerator.
• Standard P calibration solution (SPCS), or working standards, 2.0, 1.6, 1.2, 0.8, 0.4, and
0.0 mg P L-1. Make fresh weekly. Store in the refrigerator. Allow to equilibrate to room
temperature before use. To six 50-mL volumetric flasks, add as follows:
o 2.0 mg P L'1= 25 mL WSSPS
o 1.6 mg P L'1 = 20 mL WSSPS
o 1.2mgPL'1 = 15 mL WSSPS
o 0.8 mg P L'1 = 10 mL WSSPS
o 0.4 mg P L'1 = 5 mL WSSPS
o 0.0 mg P L'1 = 0 mL WSSPS (blank)
• Dilute each SPCS to mark with extracting solution and invert to thoroughly mix.
• Quality Control Samples: 0.1 mg P L-1 solution made from SSPS; blanks; and selected
SPCS. In addition, SSL soil standard and WEPAL ISE's (Wageningen Evaluating
Programmes for Analytical Laboratories, International Soil Exchange) from the Netherlands
are routinely included in a batch for quality control.
5.19.6 Procedure
1. Weigh 1.0 g of <2mm or fine-grind, air-dry soil to the nearest mg into a 50-mL centrifuge
tube. If sample is moist, weigh enough soil to achieve ~ 1.0 g of air-dry soil.
2. Dispense 20.0 mL of extracting solution to tube.
3. Transfer the sample to the shaker. Shake for 30 min at 200 oscillations min"1 at room
temperature (20°C± 2°C).
4. Remove the sample from the shaker. Centrifuge at 2000 rpm for 10 min, decant, filter, and
collect extract in receiving cup. If extracts are not to be determined immediately after
collection, then store samples at 4°C. Analyze samples within 72 h.
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5. Use the pipette to transfer a 5-mL aliquot of the sample to a plastic cup. Also transfer a 5-
ml_ aliquot of each SPCS to a plastic cup. Use a clean pipette tip for each sample and
SPCS.
6. Dispense 5 ml_ of color developing reagent to sample aliquot and to each SPCS. Swirl to
mix. Do not place sample cups close together as carbon dioxide is released and solution will
bubble. The color reaction requires a minimum of 20 min before analyst records readings.
Allowing 1 h for color development usually improves results. Color will remain stable for 24
h.
7. Transfer sample extract and SPCS to cuvettes.
8. Set the spectrophotometer to read at 882 nm. Autozero with calibration blank.
9. Calibrate the instrument using the SPCS. The data system will then associate the
concentrations with the instrument responses for each SPCS. Rejection criteria for SPCS, if
R2 <0.99.
10. Run samples using calibration curve. Sample concentration is calculated from the
regression equation. Rejection criteria for batch, if blanks as samples > 0.01; if SPCS as
samples > ± 20%; if SSL standard > ± 20% mean; and if ISE > (3 x MAD), where MAD =
median of absolute deviations. Record results to the nearest 0.01 unit for the sample extract
and each SPCS.
11. If samples are outside calibration range, dilute sample extracts with extracting solution and
re-analyze.
5.19.7 Calculation
Convert the extract P (mg L"1) to soil P (mg kg ~1) as follows:
Soil P (mg kg'1) = [(AxBxCxRx 1000)/E]
where:
A = Sample extract reading (mg L"1)
B = Extract volume (L)
C = Dilution, if performed
R = Air-dry/oven-dry ratio (procedure 3D1) or field-moist/oven-dry ratio
(procedure 3D2)
1000 = Conversion factor to kg-basis
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E = Sample weight (g)
5.19.8 Reporting
Report data to the nearest 0.1 mg P kg"1 soil.
5.19.9 References
Buurman, P, B. van Lagen, and E.J. Velthorst. 1996. Manual for soil and water analysis.
Backhuys Publ., Leiden, the Netherlands.
Burt, R., M..D. Mays, E.G. Benham, and M.A. Wilson. 2002. Phosphorus characterization and
correlation with properties of selected benchmark soils of the United States. Commun.
Soil Sci. Plant Anal. 33:117-141.
5.20 Phosphorus (Mehlich No. 3)
5.20.1 Summary of Method
A 2.5-g soil sample is shaken with 25 ml_ of Mehlich No. 3 extracting solution for 5 min. The
sample is centrifuged until solution is free of soil mineral particles, and then filtered until clear
extracts are obtained. Dilute 0.5-mL of sample extract with 13.5-mL of working solution.
Absorbance of the solution is read using a spectrophotometer at 882 nm. Data are reported as
mg P kg"1 soil.
5.20.2 Interferences
The Mo blue methods, which are very sensitive for P, are based on the principle that in an acid
molybdate solution containing orthophosphate ions, a phosphomolybdate complex forms that
can be reduced by ascorbic acid, SnCI2, and other reducing agents to a Mo color, with ascorbic
acid most commonly used by agricultural laboratories (Murphy and Riley, 1962). The intensity of
blue color varies with the P concentration but is also affected by other factors such as acidity,
arsenates, silicates, and substances that influence the oxidation-reduction conditions of the
system (Olsen and Sommers, 1982).
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5.20.3 Precautions
Wear protective clothing (coats, aprons, sleeve guards, and gloves) and eye protection (face
shields, goggles, or safety glasses). When preparing reagents, exercise special care. Many
metal salts are extremely toxic and may be fatal if ingested. Thoroughly wash hands after
handling these metal salts. Restrict the use of concentrated H2SO4 and HCI to a fume hood.
Use safety showers and eyewash stations to dilute spilled acids and bases. Use sodium
bicarbonate and water to neutralize and dilute spilled acids.
5.20.4 Equipment/Materials
Electronic balance, ±1.0-mg sensitivity
Mechanical reciprocating shaker, 200 oscillations min-1, 1 1/2 in strokes, Eberbach 6000,
Eberbach Corp., Ann Arbor, Ml
Centrifuge tubes, 50-mL, polyethylene
Funnel, 60° angle, long stem, 50-mm diameter
Filter paper, Whatman 42, 150 mm
Centrifuge, Centra, GP-8, Thermo I EC, Needham Heights, MA
Pipettes, electronic digital, 1000 uL and 10 ml_, with tips, 1000 uL and 10 mL
Dispenser, 30 mL or 10 mL
Cups, plastic
Cuvettes, plastic, 4.5-mL, 1-cm light path, Daigger Scientific
Spectrophotometer, UV-Visible, Dual-View, Varian, Gary 50 Cone, Varian Australia Pty Ltd.
Computer with Gary WinUV software, Varian Australia Pty Ltd., and printer
5.20.5 Reagents
• Reverse osmosis deionized (RODI) water, ASTM Type I grade of reagent water
• Sulfuric acid (H2SO4), concentrated, 36 N, trace pure grade
Mehlich No. 3 Extracting solution (0.2 N CH3COOH; 0.25 N NH4NO3; 0.015 N NH4F; 0.13
N HNO3; 0.001 M EDTA). Premixed Mehlich No. 3 Extractant, Special-20, Hawk Creek
Laboratory, Rural Route 1, Box 686, Simpson Road, Glen Rock, PA, 17327.
• Sulfuric-tartarate-molybdate solution (STMS). Dissolve 100 g of ammonium molybdate
tetrahydrate [(NH4)6Mo7O24'4H2O] in 500 mL of RODI water. Dissolve 2.425 g of antimony
potassium tartarate (potassium antimony tartarate hemihydrate. [K(SbO)C4H4O6*1/2H2O] in
the ammonium molybdate solution. Slowly and carefully add 1400 mL of concentrated
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H2SO4 and mix well. Cool and dilute to 2 L with RODI water. Store in the dark in the
refrigerator.
• Ascorbic acid solution. Dissolve 8.8 g of ascorbic acid in RODI water and dilute to 100- mL
with RODI water. Make fresh daily.
• Working ascorbic acid molybdate solution (WAMS). Dilute 20 ml_ of STMS solution and 10
ml_ of the ascorbic acid solution with RODI water to make 1-L. Allow solution to come to
room temperature before using. Prepare fresh daily.
Working stock standard P solution (WSSPS), 100.0 mg P L-1. In a 1-L volumetric flask,
dissolve 0.4394 g primary standard grade anhydrous potassium dihydrogen phosphate
(KH2PO4) (dried for 2 h at 110°C) in about 800 ml_ extracting solution. Dilute to 1-L volume
with extracting solution and invert to thoroughly mix. Store in polyethylene containers.
Make fresh weekly. Store in a refrigerator.
Standard P calibration solutions (SPCS) or working standards, 12.0, 10.0, 8.0, 4.0, 2.0,
1.0, 0.8, 0.4 and 0.0 mg P L-1. Prepare fresh weekly. Store in a refrigerator. Allow to
equilibrate to room temperature before use. In nine 250-mL volumetric flasks add as
follows:
o 12.0 mg P L'1 = 30 mL WSSPS
o 10.0mg PL'1 = 25 mL WSSPS
o 8.0 mg P L'1 = 20 mL WSSPS
o 4.0 mg P L'1 = 10 mL WSSPS
o 2.0 mg P L'1 = 5 mL WSSPS
o 1.0 mg PL'1 = 2.5 mL WSSPS
o 0.8 mg P L'1 = 2 mL WSSPS
o 0.4 mg P L-1 = 1 mL WSSPS
o 0.0 mg P L1 = 0 mL WSSPS (blank)
• Dilute each SPCS to the mark with extracting solution and invert to thoroughly mix.
• Quality Control Samples: 0.1 mg P L -1 solution made from SSPS; blanks; and selected
SPCS. In addition, SSL soil standard and WEPAL ISE's (Wageningen Evaluating
Programmes for Analytical Laboratories, International Soil Exchange) from The
Netherlands are routinely included in a batch for quality control.
5.20.6 Procedure
1. Weigh 2.5 g of <2-mm or fine-grind, air-dry soil to the nearest mg into a 50-mL centrifuge
tube. If sample is moist, weigh enough soil to achieve ~ 2.5 g of air-dry soil.
2. Dispense 25.0 mL of extracting solution to the tube.
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3. Transfer the sample to the shaker. Shake for 5 min at 200 oscillations min-1 at room
temperature (20°C± 2°C).
4. Remove the sample from the shaker. Centrifuge at 2000 rpm for 10 min, decant, filter, and
collect extract in receiving cups. If extracts are not to be determined immediately after
collection, then store samples at 4°C. Analyze samples within 72 h.
5. Use the pipette to transfer a 1 ml_ (or 0.5-mL) aliquot of the sample to a plastic cup. Also
transfer a 1 ml_ (or 0.5-mL) aliquot of each SPCS to a plastic cup. Use a clean pipette tip for
each sample and SPCS.
6. Dispense 27 mL (or 13.5 mL) of the WAMS to sample aliquot and to each SPCS. Swirl to
mix. The color reaction requires a minimum of 20 min before analyst records readings.
Allowing 1 h for color development usually improves results. Color will remain stable for 6 h.
7. Transfer sample extract and SPCS to cuvettes.
8. Set the spectrophotometer to read at 882 nm. Autozero with calibration blank.
9. Calibrate the instrument by using the SPCS. The data system will then associate the
concentrations with the instrument responses for each SPCS. Rejection criteria for SPCS, if
R2 <0.99.
10. Run samples using calibration curve. Sample concentration is calculated from the
regression equation. Rejection criteria for batch, if blanks as samples > 0.01; if SPCS as
samples > ± 20%; if SSL standard > ± 20% mean; and if ISE > (3 x MAD), where MAD =
median of absolute deviations. Record results to the nearest 0.01 unit for the sample extract
and each SPCS.
11. If samples are outside calibration range, dilute sample extracts with extracting solution and
re-analyze.
5.20.7 Calculation
Convert extract P (mg L"1) to soil P (mg kg"1) as follows:
Soil P (mg kg ~1) = [(A x B x C x R x 1000)/E]
where:
A = Sample extract reading (mg L"1)
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B = Extract volume (L)
C = Dilution, if performed
5.20.8 Reporting
Report data to the nearest 0.1 mg P kg 1 soil.
5.20.9 References
Mehlich, A. 1984. Mehlich 3 soil text extractant: A modification of Mehlich 2 extractant.
Commun. Soil Sci. Plant Anal. 15:1409-1416.
Murphy, J., and J.R. Riley. 1962. A modified single solution method for the determination of
phosphate in natural waters. Anal. Chem. Acta. 27:31-36.
Olsen, S.R., and L.E. Sommers. 1982. Phosphorus, p. 403-430. In A.L Page, R.H. Miller, and
D.R. Keeney (eds.) Methods of soil analysis. Part 2. Chemical and microbiological
properties. 2nd ed. Agron. Monogr. 9. ASA and SSSA, Madison, Wl.
Soil and Plant Analysis Council, Inc. 1999. Handbook on reference methods for soil analysis.
Council on Soil Testing and Plant Analysis. CRC Press, Boca Raton, FL.
Iran, T.S., and R.R. Simard. 1993. Mehlich Ill-extractable elements, p. 43-50. In M.R. Carter
(ed.) Soil sampling and methods of analysis. Can. Soc. Soil Sci. Lewis Publ., Boca
Raton, FL.
5.21 Trace Elements (Ag, Ba, Be, Cd, Co, Cr, Cu, Hg, Mn, Mo, Ni, P,
Pb, Sb, Se, Sn, Sr, Tl, V, W, and Zn)
5.21.1 Summary of Method
The approach of this digestion methodology is to maximize the extractable concentration of
elements in digested soils while minimizing the matrix interferences such as found in digestion
procedures that use HF acid. This method (4H1a1) follows EPA Method 3051A. A 500-mg <2-
mm soil separate which has been air-dried and ground to < 200 mesh (75 |im) is weighed into a
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100-ml Teflon (PFA) sample digestion vessel. To the vessel, 9.0 ml_ HNO3 and 3.0 ml_ HCI are
added. The vessel is inserted into a protection shield, covered and placed into a rotor with
temperature control. Following microwave digestion the rotor and samples are allowed to cool,
before quantitatively transferring the digestate into a 50-ml glass volumetric with high purity
deionized reverse osmosis water (DIRO). The samples are then allowed to cool to room
temperature before bringing to volume. Once brought to volume the samples are thoroughly
mixed and allowed to settle overnight before decanting into 50ml Falcon tubes prior to analysis.
The concentration of Ag, As, Ba, Be, Cd, Co, Cr, Cu, Hg, Mn, Mo, Ni, P, Pb, Sb, Se, Sn, V, W,
and Zn are determined using an inductively coupled plasma mass spectrometer (ICP-MS).
5.21.2 Interferences
Organic constituents may contain metals and are difficult to digest if present in high
concentrations. Certain elements are subject to volatile losses during digestion and transfer.
Certain soil minerals (e.g., quartz, feldspars) are not soluble in HNO3 + HCI.
Interferences are corrected or minimized by using an internal standard along with gas reaction
and collision cell and technology. Also, careful selection of specific masses for data reporting is
important. Interference corrections are made by ICP software. Samples and standards are
matrix-matched to help reduce interferences.
5.21.3 Precautions
Wear protective clothing and eye protection. When preparing reagents, exercise special care.
Restrict the use of concentrated acids to the fume hood. Wash hands thoroughly after handling
reagents. Filling the digestion vessel to greater than 25 percent of the free volume or adding
organic reagents or oxidizing agents to the vessel may result in explosion of the digestion
microwave system.
5.21.4 Equipment/Materials
• Electronic balance, ±1.0-mg sensitivity
• Pipet(s) capable of delivering 3 and 9 ml_, Omnifit Corp. manufacturers variable volume,
(10-ml maximum) pipettes suitable for HNO3 and HCI delivery from 2.5 L bottles
• Volumetric flasks, class A glass, 50 ml_
• Polypropylene bottles, 60 ml_, with cap
• Electronic balance (±0.1 mg sensitivity)
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• Microwave oven, CEM Mars 5, 14 position-HP500 Plus vessel and rotor (vessels
composed of PFA, sleeves composed of advanced composite)
• Volumetrics, 500, 250, and 50-mL class A glass
• Containers, 500-mL, polypropylene, with screw caps
• Pipettes, electronic digital, 1000 uL and 10 ml_, Rainin Instrument Co., Woburn, MA
• Inductively coupled plasma mass spectrometer (ICP-MS), Agilent 7500cx, Agilent
Technologies Inc. Wilmington, Delaware
• Computer, with ICP-MS ChemStation software ver. B.03.07, Agilent Technologies Inc.
Wilmington, Delaware
• Heat Exchanger, G1879B, Agilent Technolgies
• Compressed gasses, argon (minimum purity = 99.99%), hydrogen (minimum purity =
99.999%) and helium (minimum purity = 99.999%)
• Autosampler, ASX-500 Series, Agilent Technolgies Inc. Wilmington, De.
Quartz torch, for use with HMI, Part No. G3270-80027
• Peristaltic pump (for automatic injection of internal standard)
5.21.5 Reagents
• Deinonized reverse osmosis (DIRO) water, ASTM Type I grade of reagent water
• Concentrated hydrochloric acid (HCI), 12 N, trace pure grade
• Concentrated nitric acid (HNO3), 16 N, trace pure grade
• Primary standards: 1000 mg L-1, from High Purity Standards, Charleston, SC. Single and
mixed element standards are manufactured in dilute HNO3, HNO3 + HF, or H2O.
5.21.6 Procedure
Microwave Acid Digestion
1. About 500 mg of fine-earth (<2-mm) or a specific particle size separate ground to <200-
mesh (75 urn) is weighed to the nearest 0.1 mg in a 100-mL digestion vessel.
Note: If sample is principally composed of organic materials (organic C > 15%), perform a
preliminary digestion in the muffle furnace in an digestion crucible: 250°C for 15 min, 450°C for
15 min, followed by 550°C for 1 h.
2. Pipet 9.0 ml_ HNO3 and 3.0 ml_ HCI into the sample and allow to completely wet. Add acids
in the fume hood. Allow acids to react and vent in uncovered vessels for about 30 min.
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3. Place covered vessels in protective sleeve, cover and place into rotor, attach pressure
sensor to reference vessel, tighten all vessels to proper torque.
4. Place digestion rotor in the microwave oven and insert the temperature probe into the
reference vessel. Attach the temperature and pressure sensor cables to the fittings in the
microwave.
5. Microwave settings are as follows:
• 1200 watts at 100% power for 5.5 min until 175°C
• Hold at 175°C for 4.5 min
• Cool for 5 min
6. After cooling, disconnect temperature probe and pressure sensor from microwave.
7. Remove rotor from oven, and place in fume hood.
8. Open each vessel carefully and then quantitatively transfer contents of vessel to a 50ml_
volumetric flask with DIRO water.
9. Cap flask and mix well by inverting. Allow samples to cool to room temperature. When
cool, fill samples to volume with DIRO water and let settle overnight.
10. Decant contents into a labeled 60-mL polypropylene container.
11. Prepare working standards of a blank, reference soil sample from the SSL repository, NIST
or other standard reference material and blank by the same digestion method. Run two of
these standards and a blank with each set of 14 samples.
ICP-MS Calibration Standards, Set-Up, and Operation
12. Trace Method Stock C - Commercially prepared solution containing 10ug/ml_ Sb, As, Ba,
Be, Cd, Cr, Co, Cu, Pb, Mo, Ni, Se, Sn, W, V, Zn, 1ug/ml_ Ag
13. Trace Method Stock A - Commercially prepared solution containing10ug/ml_ Mn, P, Sr
14. Mercury Stock Standard - Commercially prepared solution containing1ug/ml_ Hg
15. Standard Matrix - Place 500ml_ DIRO in a 1 L-volumetric flask, add 0.250ml_ 1000ug/ml_ Au
and 18ml_ concentrated nitric acid. Fill to volume with DIRO and mix well.
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16. TM7 - In a 1L flask add 300ml DIRO, 18ml_ concentrated nitric acid, 6ml_ concentrated
hydrochloric acid, 0.250ml of 1000|jg/mL Au, 1.0ml_ of Mercury Stock standard (1.13.6.15),
fill to volume with DIRO and mix well.
17. TM6 - In a 1L flask add 300ml DIRO, 18ml_ concentrated nitric acid, 6ml_ concentrated
hydrochloric acid, 0.250ml of 1000ug/ml_ Au, 0.5ml_ of Mercury Stock standard (1.13.6.15),
fill to volume with DIRO and mix well.
18. TM5 - In a 500ml_ volumetric add SOOmL DIRO and 50.0ml_ Trace Method Stock A
(1.13.6.14), fill to volume with DIRO and mix well.
19. TM4- In a SOOmL volumetric add SOOmL DIRO and 12.5mL Trace Method Stock A
(1.13.6.14), fill to volume with DIRO and mix well.
20. TM3 - In a SOOmL volumetric flask, add SOOmL DIRO, 9mL of concentrated nitric acid,
0.125ml of 1000ug/mLAu and O.SmL Trace Method Stock C (1.13.6.15). Fill to volume with
DIRO, mix well.
21. TM2 - In a SOOmL volumetric flask add SOOmL of Standard Matrix (1.13.6.16) and 50mL TM3
(1.13.6.15). Fill to volume with Standard Matrix (1.13.6.16), mix well.
22. TM1 - In a SOOmL volumetric add SOOmL of Standard Matrix (1.13.6.16) and 50mLTM2
(1.13.6.22). Fill to volume with Standard Matrix (1.13.6.16), mix well.
23. TMO - In a SOOmL volumetric flask add SOOmL DIRO, 9mL concentrated nitric acid, 3mL
concentrated hydrochloric acid, 0.125ml of 1000ug/mL Au, fill to volume with DIRO and mix
well.
24. Internal Standard (1ug/mL Li6 , Sc, Ge, Y, In, Tb, Bi) - In a 1L flask add SOOmL DIRO, 18mL
concentrated nitric acid, 6mL concentrated hydrochloric acid, .250ml of 1000ug/mL Au and
100mL of commercially prepared Internal Standard Mix, fill to volume with DIRO and mix
well.
25. Rinse - In 2L flask add SOOmL DIRO, 58mL concentrated nitric acid, fill to volume with DIRO
and mix well.
26. Rinse #1 - In a 1L flask add SOOmL DIRO, 29mL concentrated nitric acid, 1ml of 1000ug/mL
Au, fill to volume with DIRO and mix well.
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27. Rinse #2 - In a 1L flask add SOOmL DIRO, 15ml_ concentrated nitric acid, 45ml_
concentrated hydrochloric acid and 1ml of 1000|jg/mL Au, fill to volume with DIRO and mix
well.
28. Sample diluent - In a 1L flask add SOOmL DIRO, 0.277ml_ of 1000ug/ml_ Au, fill to volume
with DIRO and mix well.
29. Standard concentrations in ug/mL for each element.
Element
Sb
As
Ba
Be
Cd
Cr
Co
Cu
Pb
Mn
Hg
Mo
Ni
P
Se
Ag
Sr
Sn
W
V
Zn
TMO
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
TM1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.01
0.1
0.1
0.1
0.1
TM2
1
1
1
1
1
1
1
1
1
1
1
1
0.1
1
1
1
1
TM3
10
10
10
10
10
10
10
10
10
10
10
10
10
1
10
10
10
10
TM4
250
250
250
TM5
1000
1000
1000
TM6
0.5
TM7
1
30. Reporting m/z and tune step for each element analyzed.
Element
Sb
As
Ba
m/z
121
75
137
Tune 1
(H2)
Tune 2
(He)
X
Tune 3
(No
Gas)
X
X
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Element
Be
Cd
Cr
Co
Cu
Pb
Mn
Hg
Mo
Ni
P
Se
Ag
Sr
Sn
W
V
Zn
m/z
9
111
52
59
63
208
55
202
98
60
31
78
107
88
118
182
51
66
Tune 1
(H2)
X
X
Tune 2
(He)
X
X
X
X
X
X
X
X
X
X
X
X
Tune 3
(No
Gas)
X
X
X
X
X
X
31. Use the ICP-MS with a micromist nebulizer, quartz torch for micro flow nebulizer and quartz
spray chamber to analyze samples. Internal standard is added via peristaltic pump using
0.19 mm id. pump tubing. Internal standard and samples or standards are mixed via coil
prior to entering the nebulizer. Samples are diluted 1:10 or greater as necessary prior to
analysis with sample diluent (1.13.6.29). Perform instrument checks (tune for sensitivity,
resolution axis, P/A factor, internal standard RSD, torch alignment, EM tune) prior to
analysis as discussed in operation manual of instrument. Check instrument gas pressures to
ensure pressures are correct and of adequate supply.
32. Typical tune values for trace analysis method.
Tune 1 (H2)
Plasma Parameters
RF Power
RF Matching
Smpl Depth
Torch-H
Torch-V
Carrier Gas
Makeup Gas
1550W
1.78V
8.5mm
0.4mm
0.2mm
0.90L/min
0.15L/min
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Optional Gas 0.0
Nebulizer Pump O.IOrps
Sample Pump 0.0
S/C Temp 2degC
Ion Lenses
Extract 1
Extract 2
Omega Bias-ce
Omega Lens-ce
Cell Entrance
QP Focus
Cell Exit
Q-Pole Parameters
AMU Gain
AMU Offset
Axis Gain
Axis Offset
QP Bias
Octapole Parameters
OctP RF
OctP Bias
Reaction Cell
Reaction Mode
H2Gas
He Gas
Optional Gas
Detector Parameters
Discriminator
Analog HV
Pulse HV
0.0V
-136.0V
-28V
0.0V
-40V
-8V
-40V
130
124
0.9996
0.04
-15.0V
150V
-18.0V
ON
2.5ml_/min
O.OmL/min
S.OmV
1690V
1260V
Tune 2 (He)
Plasma Parameters
RF Power 1550W
RF Matching 1.78V
Smpl Depth 8.5mm
Torch-H 0.4mm
Torch-V 0.2mm
Carrier Gas 0.90L/min
Makeup Gas 0.15L/min
Optional Gas 0.0
Nebulizer Pump O.IOrps
Sample Pump 0.0
S/C Temp 2degC
Ion Lenses
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Extract 1
Extract 2
Omega Bias-ce
Omega Lens-ce
Cell Entrance
QP Focus
Cell Exit
Q-Pole Parameters
AMU Gain
AMU Offset
Axis Gain
Axis Offset
QP Bias
Octapole Parameters
OctP RF
OctP Bias
Reaction Cell
Reaction Mode
H2Gas
He Gas
Optional Gas
Detector Parameters
Discriminator
Analog HV
Pulse HV
0.0V
-136.0V
-28V
0.0V
-40V
-10V
-40V
130
124
0.9996
0.04
-16.0V
150V
-18.0V
ON
O.OmL/min
6.0ml_/min
S.OmV
1690V
1260V
Tune 3 (No Gas)
Plasma Parameters
RF Power 1550W
RF Matching 1.78V
Smpl Depth 8.5mm
Torch-H 0.4mm
Torch-V 0.2mm
Carrier Gas 0.90L/min
Makeup Gas 0.15L/min
Optional Gas 0.0
Nebulizer Pump O.IOrps
Sample Pump 0.0
S/C Temp 2degC
Ion Lenses
Extract 1 0.0V
Extract 2 -136.0V
Omega Bias-ce -28V
Omega Lens-ce 0.0V
Cell Entrance -30V
QP Focus 3V
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Cell Exit -30V
Q-Pole Parameters
AMU Gain 130
AMU Offset 124
Axis Gain 0.9996
Axis Offset 0.04
QP Bias -3.0V
Octapole Parameters
OctP RF 150V
OctP Bias -6.0V
Reaction Cell
Reaction Mode OFF
H2Gas
He Gas
Optional Gas
Detector Parameters
Discriminator S.OmV
Analog HV 1690V
Pulse HV 1260V
33. Use the blank standard solution to dilute those samples with concentrations greater than the
high standard. Rerun all elements and use only the data needed from the diluted analysis.
34. Establish detection limits using the blank standard solution. The instrumental detection
limits are calculated by using 3 times the standard deviation of 10 readings of the blank.
These values establish the lower detection limits for each element. Analyzed values lower
than the detection limits are reported as "ND" or non-detected.
5.21.7 Calculation
The calculation of mg kg"1 of an element in the soil from ug L"1 in solution is as follows:
Analyte concentration in soil (mg kg ) = [AxBxCxRx 1000]/E x 1000
Where:
A = Sample extract reading (ug L"1)
B = Extract volume (L)
C = Dilution, if performed
R = Air-dry/oven-dry ratio (procedure 3D1)
1000 = Conversion factor in numerator to kg-basis
E = Sample weight (g)
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1000 = Factor in denominator (|jg mg"1)
5.21.8 Reporting
Analysis is generally done on one mass per element, if more than one mass is analyzed only
the reporting mass is used for data reporting purposes. The particle-size fraction digested needs
to be identified with each sample. Data are reported to the nearest 0.01 mg kg"1.
5.21.9 References
Burt, R., M.A. Wilson, T.J. Keck, B.D. Dougherty, D.E. Strom, and J.A. Lindhal. 2002. Trace
element speciation in selected smelter-contaminated soils in Anaconda and Deer Lodge
Valley, Montana, USA. Adv. Environ. Res. 8:51-67.
Burt, R., M.A. Wilson, M.D. Mays, and C.W. Lee. 2003. Major and trace elements of selected
pedons in the USA. J. Environ. Qual. 32:2109-2121.
Gambrell, R.P. 1994. Trace and toxic metals in wetlands-A review. J. Environ. Qual. 23:883-
891.
Holmgren, G.G.S., M.W. Meyer, R.L. Chaney, and R.B. Daniels. 1993. Cadmium, lead, zinc,
copper, and nickel in agricultural soils of the United States of America. J. Environ. Qual.
22:335-348.
Jersak, J., R. Amundson, and G. Brimhall, Jr. 1997. Trace metal geochemistry in Spodosols of
the Northeastern United States. J. Environ. Qual. 26:551-521.
Keller, C., and J.C. Vedy. 1994. Distribution of copper and cadmium fractions in two forest
soils. J. Environ. Qual. 23:987-999.
Pierzynski, G.M., and A. P. Schwab. 1993. Bioavailability of zinc, cadmium, and lead in a
metal-contaminated alluvial soil. J. Environ. Qual. 22:247-254.
Tiller, K.G. 1989. Heavy metals in soils and their environmental significance. In B.A. Stewart
(ed.)Adv. SoilSci. 9:113-142.
Wilcke, W., and W. Amelung. 1996. Small-scale heterogeneity of aluminum and heavy metals
in aggregates along a climatic transect. Soil Sci. Soc. Am. Proc. 60:1490-1495.
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Wilcke, W., S. Muller, N. Kanchanakool, and W. Zech. 1998. Urban soil contamination in
Bangkok: Heavy metal and aluminum partitioning in topsoils. Geoderma 86:211-228.
5.22 Bulk Density (Hammered Core, Field State)
Density is defined as mass per unit volume. Soil bulk density of a sample is the ratio of the
mass of solids to the total or bulk volume. This total volume includes the volume of both solids
and pore space. Bulk density is distinguished from particle density which is mass per unit
volume of only the solid phase. Particle density excludes pore spaces between particles. As
bulk density (Db) is usually reported for the <2-mm soil fabric, the mass and volume of rock
fragments are subtracted from the total mass and volume.
Bulk density is highly dependent on soil conditions at the time of sampling. Changes in soil
volume due to changes in water content will alter bulk density. Soil mass remains fixed, but the
volume of soil may change as water content changes (Blake and Hartge, 1986). Bulk density, as
a soil characteristic, is actually a function rather than a single value. Therefore, subscripts are
added to the bulk density notation, Db, to designate the water state of the sample when the
volume was measured. The SSL uses the bulk density notations of Dbf, Db33, and Dbod for field-
state, 33-kPa equilibration, and oven-dry, respectively.
Field-state (Dbf) is the bulk density of a soil sample at field-soil water content at time of
sampling. The 33-kPa equilibration (Db33) is the bulk density of a soil sample that has been
desorbed to 33kPa (1/3 bar). The oven-dry (Dbod) is the bulk density of a soil sample that has
been dried in an oven at 110°C. Bulk density also may be determined for field-moist soil cores of
known volume. The bulk density of a weak or loose soil material for which the clod or core
method is unsuitable may be determined by the compliant cavity method.
5.22.1 Summary of Method
A metal cylinder is pressed or driven into the soil. The cylinder is removed extracting a sample
of known volume. The moist sample weight is recorded. The sample is then dried in an oven
and weighed.
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5.22.2 Interferences
During coring process, compaction of the sample is a common problem. Compression can be
observed by comparing the soil elevation inside the cylinder with the original soil surface outside
the cylinder. If compression is excessive, soil core may not be a valid sample for analysis. Rock
fragments in the soil interfere with core collection. Dry or hard soils often shatter when
hammering the cylinder into the soil. Pressing the cylinder into the soil reduces the risk of
shattering the sample.
If soil cracks are present, select the sampling area so that crack space is representative of
sample, if possible. If this is not possible, make measurements between the cracks and
determine the aerial percentage of total cracks or of cracks in specimen.
5.22.3 Equipment
• Containers, air-tight, tared, with lids
• Electronic balance, ±0.01-g sensitivity
Oven110°C
• Sieve, No. 10 (2 mm-openings)
• Coring equipment. Sources described in Grossman and Reinsch (2002).
5.22.4 Procedure
1. Record the empty core weights (CW).
2. Prepare a flat surface, either horizontal or vertical, at the required depth in sampling pit.
3. Press or drive core sampler into soil. Use caution to prevent compaction. Remove core from
the inner liner, trim protruding soil flush with ends of cylinder, and place in air-tight container
for transport to laboratory. If soil is too loose to remain in the liner, use core sampler without
the inner liner and deposit only the soil sample in air-tight container. Moisture can also be
pushed directly into a prepared face. For fibrous organic materials, trim sample to fit snugly
into a can.
4. Dry core in an oven at 110°C until weight is constant. Record oven-dry weight (ODW).
5. Measure and record cylinder volume (CV).
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6. If sample contains rock fragments, wet sieve sample through a 2-mm sieve. Dry and weigh
the rock fragments that are retained on sieve. Record weight of rock fragments (RF).
7. Determine density of rock fragments (PD).
5.22.5 Calculations
Db = (ODW - RF - CW)/[CV - (RF/PD)]
where:
Db = Bulk density of < 2-mm fabric at sampled, field water state (g cm-3)
ODW = Oven-dry weight
RF = Weight of rock fragments
CW = Empty core weight
CV = Core volume
PD = Density of rock fragments
5.22.6 Report
Bulk density is reported as g cc-1 to the nearest 0.01 g cm"3.
5.22.7 References
Grossman, R.B. and T.G. Reinsch. 2002. Bulk density and linear extensibility, p. 201-228. In
J.H. Dane and G.C. Topp (eds.) Methods of soil analysis, Part 4. Physical methods. Soil
Sci. Am. Book Series No. 5. ASA and SSSA, Madison, Wl.
5.23 Bulk Density (Saran Clod, Field State)
Density is defined as mass per unit volume. Soil bulk density of a sample is the ratio of the
mass of solids to the total or bulk volume. This total volume includes the volume of both solids
and pore space. Bulk density is distinguished from particle density which is mass per unit
volume of only the solid phase. Particle density excludes pore spaces between particles. As
bulk density (Db) is usually reported for the <2-mm soil fabric, the mass and volume of rock
fragments are subtracted from the total mass and volume.
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Bulk density is highly dependent on soil conditions at the time of sampling. Changes in soil
volume due to changes in water content will alter bulk density. Soil mass remains fixed, but the
volume of soil may change as water content changes (Blake and Hartge, 1986). Bulk density, as
a soil characteristic, is actually a function rather than a single value. Therefore, subscripts are
added to the bulk density notation, Db, to designate the water state of the sample when the
volume was measured. The SSL uses the bulk density notations of Dbf, Db33, and Dbod for field-
state, 33-kPa equilibration, and oven-dry, respectively.
Field-state (Dbf) is the bulk density of a soil sample at field-soil water content at time of
sampling. The 33-kPa equilibration (Db33) is the bulk density of a soil sample that has been
desorbed to 33kPa (1/3 bar). The oven-dry (Dbod) is the bulk density of a soil sample that has
been dried in an oven at 110°C. Bulk density also may be determined for field-moist soil cores
of known volume (procedure 3B6a). The bulk density of a weak or loose soil material for which
the clod or core method is unsuitable may be determined by the compliant cavity method
(procedure 3B3a).
5.23.1 Summary of Method
Field-occurring fabric (clods) is collected from the face of an excavation. One coat of plastic
lacquer is applied in the field. Additional coats of plastic lacquer are applied in the laboratory.
In its field-water state or after equilibration, the clod is weighed in air to measure its mass and in
water to measure its volume. After the clod is dried in oven at 110°C, its mass and volume are
determined again. A correction is made for the mass and volume of rock fragments and plastic
coatings (Brasher et al., 1966; Blake and Hartge, 1986; Grossman and Reinsch, 2002).
5.23.2 Interferences
Errors are caused by non-representative samples. Only field-occurring fabric (clods) should be
sampled. The whole soil bulk density may be overestimated because sampled clods frequently
exclude the crack space between clods (Grossman and Reinsch, 2002).
The penetration of plastic lacquer into the voids of sandy or organic soils interferes with the
corrections for mass and volume of the plastic coat and with the accuracy of water content
determinations. Penetration can be reduced by spraying water on the clod followed by
immediately dipping the clod in the plastic lacquer.
Loss of soil during the procedure will void the analyses because all calculations are based on
the oven-dry soil mass. Holes in the plastic coating, which are detected by escaping air bubbles
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from submerged clod, introduce errors in volume measurement. An inadequate evaporation of
the plastic solvent results in overestimation of the soil mass. A drying time of 1 h is usually
sufficient time for evaporation of solvent. However, clods with high organic matter content may
need to dry longer.
Bulk density is reported for <2-mm soil fabric. Correction for rock fragments >2-mm requires
either knowledge or assumption of the rock fragment density. Estimate or measurement errors
of rock fragment density will affect the accuracy of the value for soil bulk density. Rock
fragments may contain water which complicates the application to actual water-holding capacity.
5.23.3 Precautions
Methyl ethyl ketone (MEK) is extremely flammable. A type B fire extinguisher should be in close
proximity in the laboratory. No open flames or nearby operation of electrical equipment are
permitted while using MEK. The MEK vapor is classified as a sensory irritant. The 8-h time-
weighted average (TWA) exposure limit is 200 ppm, and the short-term exposure limit (STEL) is
300 ppm (Occupational Safety and Health Administration, 1989). Avoid physical contact. Use
with adequate ventilation. In closed buildings, use a fume hood. Keep in tightly closed
containers. Use safety glasses, proper gloves, and a lab coat. Wash hands immediately after
handling MEK. Additional information on the safe handling of MEK is available in Chemical
Safety Data Sheet SD-83, Manufacturing Chemists' Association, Inc., 1825 Connecticut Ave.
NW, Washington, DC.
Saran F-310 resin will decompose rapidly at temperatures >200°C releasing hydrogen chloride
gas. Avoid contact with Fe, Zn, Cu, and Al in solution. Avoid all contact with strong bases.
5.23.4 Equipment
• Electronic balance, ±0.01-g sensitivity
• Rigid shipping containers. The SSL uses a corrugated box with compartments.
• Plastic bags, 1 ml_, 127 x 89 x 330 mm
• Wire. The SSL uses a 28-awg coated copper wire.
• Hairnets
• Stock tags, 25.4-mm (1-in) diameter paper tag, with metal rim
• Hook assembly for weighing below balance
• Plexiglass water tank mounted on a fulcrum and lever to elevate tank
Oven, 110°C
• Sieve, No. 10 (2 mm openings)
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• Rope, 3 m
• Clothespins
• Silt loam soil
• Hot plate
• Spray bottle
• Liquid vapor trap. The SSL constructs a tin enclosure over hot plate with a chimney and
duct to transfer vapor to water stream.
5.23.5 Reagents
• Methyl ethyl ketone (MEK), practical (2-butanone)
• Water
• Alcohol
• Liquid detergent. The SSL uses Liqui Nox.
• Dow Saran F-310 Resin, available from Dow Chemical Company.
• Plastic lacquer. Prepare plastic lacquer with resin to solvent ratios of 1:4 and 1:7 on a
weight basis. Fill a 3.8-L (1-gal) metal paint can with 2700 ± 200 mL of solvent. Fill to the
bottom of handle rivet. Add 540 or 305 g of resin to make 1:4 or 1:7 plastic lacquer,
respectively. For the initial field and laboratory coatings, use the 1:4 plastic lacquer. Use
1:7 plastic lacquer for the last two laboratory coats. The 1:7 plastic lacquer is used to
conserve the resin and to reduce cost. In the field, mix solvent with a wooden stick. In the
laboratory, stir solvent with a non-sparking, high speed stirrer while slowly adding resin.
Stir plastic lacquer for 15 min at 25°C. Store plastic lacquer in covered plastic or steel
containers. Acetone may be substituted for MEK.
5.23.6 Procedure
Laboratory
1. Prepare a round stock tag with sample identification number. Cut the copper wire and loop
around the clod. Record weight (TAG) of tag and wire. Loop fine copper wire around clod,
leaving a tail to which round stock tag is attached. Record weight of clod (CC1).
2. Dip clod in 1:4 plastic lacquer. Wait 7 min and then dip clod in 1:7 plastic lacquer. Wait 12
min and then dip clod in 1:7 plastic lacquer. Wait 55 min and then reweigh clod. If clod has
adsorbed >3% plastic by clod weight or smells excessively of solvent, allow longer drying
time, then reweigh clod and record weight (CC2).
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3. The clod should be waterproof and ready for volume measurement by water displacement.
Suspend the clod below the balance, submerge in water, and record weight (WMCW).
4. Dry clod in an oven at 110°C until weight is constant. Weigh oven-dry clod in air (WODC)
and in water (WODCW) and record weights.
5. If clod contains >5% rock fragments by weight, remove them from clod. Place clod in a
beaker and place on hot plate. Cover hot plate with a liquid vapor trap. Use a fume hood.
Heat clod on hot plate in excess of 200°C for 3 to 4 h. The plastic coating disintegrates at
temperatures above 200°C. After heating, clod should appear black and charred. Remove
clod from hot plate, lightly coat with liquid detergent, and add hot water.
6. Wet sieve the cool soil through a 2-mm, square-hole sieve. Dry and record weight (RF) of
rock fragments that are retained on the sieve. Determine rock fragment density by weighing
them in air to obtain their mass and in water to obtain their volume. If rock fragments are
porous and have a density similar to soil sample, do not correct clod mass and volume
measurement for rock fragments. Correct for rock fragments if these fragments can
withstand breakdown when dry soil is placed abruptly in water or calgon.
7. Correct bulk density for weight and volume of plastic coating. The coating has an air-dry
density of ~ 1.3 g cm-3. The coating loses 10 to 20% of its air-dry weight when dried in oven
at110°C.
5.23.7 Calculation
Dbf = [WODC - RF - ODPC - TAG]/{[(CC2 - WMCW)/WD] - (RF/PD) - (MPC/1.3)}
where:
Dbf = Bulk density in g cc-1 of <2 mm fabric at field-sampled water state
WODC= Weight of oven-dry coated clod
RF = Weight of rock fragments
ODPC = MPC x 0.85, weight of oven-dry plastic coat
TAG = Weight of tag and wire
CC2 = Weight of tag and wire
WMCW = Weight of coated clod in water before oven drying
WD = Water density
PD = Density of rock fragments
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MPC = [(CC2 - CC1) + FCE] x RV
where:
MPC = Weight of plastic coat before oven-drying
CC1 = Weight of clod before three laboratory plastic coats
RV = Percent estimate of remaining clod volume after cutting to obtain flat
surface (« 80%)
FCE= 1.5x[(CC2-CC1)/3]
where:
FCE = Estimate of field-applied plastic coat
Dbod = [WODC - RF - ODPC - TAG]/{[(WODC - WODCW)/WD] - (RF/PD) - (MPC/1.3)}
where:
Dbod = Bulk density in g cm-3 <2 mm fabric at oven dryness
WODCW = Weight of oven-dry coated clod in water
Wf = {[(CC2 - MPC) - (WODC - ODPC)]/[WODC - RF - ODPC - TAG]} x 100
where:
Wf = Percent water weight in sampled clod
5.23.8 Reporting
Bulk density is reported to the nearest 0.01g cm"3.
5.23.9 References
Blake, G.R. and K.H. Hartge. 1986. Bulk density, p. 363-382. In A. Klute (ed.) Methods of soil
analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9. ASA
andSSSA, Madison, Wl.
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Brasher, B.R., D.P. Franzmeier, V.T. Volassis, and S.E. Davidson. 1966. Use of saran resin to
coat natural soil clods for bulk density and water retention measurements. Soil Sci.
101:108.
Grossman, R.B., andT.G. Reinsch. 2002. Bulk density and linear extensibility, p. 202-228. In
J.H. Dane and G.C. Topp (eds.) Methods of soil analysis, Part 4. Physical methods. Soil
Sci. Am. Book Series No. 5. ASA and SSSA, Madison, Wl.
Occupational Safety and Health Administration. 1989. Air contaminants; final rule. Federal
Register. 29 CFR Part 1910. Vol. 54. No. 12. p. 2452-2453. U.S. Govt. Print. Office,
Washington, DC.
USDA 2004. Soil Survey Laboratory Methods Manual: Soil Survey Investigations Report No.
42, Version 4.0. USDA-NRCS. U.S. Govt. Print. Office, Washington, DC.
5.24 Bulk Density (Oven-Dry, clods and cores)
5.24.1 Summary of Method
Field-occurring fabric (clods or cores) is collected from the face of an excavation. One coat of
plastic lacquer is applied in the field. Additional coats of plastic lacquer are applied in the
laboratory. The clod is dried in an oven at 110°C and then weighed in air to measure its mass
and in water to measure its volume. A correction is made for the mass and volume of rock
fragments and for plastic coatings (Brasher et al., 1966; Blake and Hartge, 1986).
5.24.2 Interferences
Errors are caused by non-representative samples. Only field-occurring fabric (clods) should be
sampled. The whole soil bulk density may be overestimated because sampled clods frequently
exclude the crack space between clods (Grossman and Reinsch, 2002).
The penetration of plastic lacquer into the voids of sandy or organic soils interferes with the
corrections for mass and volume of the plastic coat and with the accuracy of water content
determinations. Penetration can be reduced by spraying water on the clod followed by
immediately dipping the clod in the plastic lacquer. Dipping should be done as quickly as
possible to reduce penetration of plastic.
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Loss of soil during the procedure will void the analyses because all calculations are based on
the oven-dry soil mass. Holes in the plastic coating, which are detected by escaping air bubbles
from submerged clod, introduce errors in volume measurement. An inadequate evaporation of
plastic solvent results in overestimation of the soil mass. A drying time of 1 h is usually
sufficient time for evaporation of solvent. However, clods with high organic matter content may
need to dry longer.
Bulk density is reported for <2-mm soil fabric. Correction for rock fragments >2-mm requires
either knowledge or assumption of the rock fragment density. Estimate or measurement errors
of rock fragment density will affect the accuracy of the value for soil bulk density. Rock
fragments may contain water, which complicates the application to actual water-holding
capacity.
5.24.3 Safety
Methyl ethyl ketone (MEK) is extremely flammable. A type B fire extinguisher should be in close
proximity in the laboratory. No open flames or nearby operation of electrical equipment are
permitted while using MEK. The MEK vapor is classified as a sensory irritant. The 8-h time
weighted average (TWA) exposure limit is 200 ppm, and the short-term exposure limit (STEL) is
300 ppm (Occupational Safety and Health Administration, 1989). Avoid physical contact. Use
with adequate ventilation. In closed buildings, use a fume hood. Keep in tightly closed
containers. Avoid physical contact. Use safety glasses, proper gloves, and a lab coat. Wash
hands immediately after handling MEK. Additional information on the safe handling of MEK is
available in Chemical Safety Data Sheet SD-83, Manufacturing Chemists' Association, Inc.,
1825 Connecticut Ave. NW, Washington, DC.
Saran F-310 resin will decompose rapidly at temperatures >200°C releasing hydrogen chloride
gas. Avoid contact with Fe, Zn, Cu, and Al in solution. Avoid all contact with strong bases.
5.24.4 Equipment
• Electronic balance, ±0.01-g sensitivity
• Rigid shipping containers. The SSL uses a corrugated box with compartments.
• Plastic bags, 1 mil, 127 x 89 x 330 mm
• Wire. The SSL uses a 28-awg coated copper wire.
• Hairnets
• Stock tags, 25.4-mm (1-in) diameter paper tag with metal rim
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• Hook assembly for weighing below balance
• Plexiglass water tank mounted on a fulcrum and lever
Oven, 110°C
• Sieve, no. 10 (2 mm openings)
• Rope, 3 m
• Clothespins
• Hot plate
• Spray bottle
• Liquid vapor trap. The SSL constructs a tin enclosure over hot plate with a chimney and
duct to transfer vapor to water stream.
• Tension table. The SSL constructs a tension table by placing porous firebricks, covered
with reinforced paper towels, in a tub of water.
5.24.5 Reagents
• Methyl ethyl ketone (MEK), practical (2-butanone).
Water.
• Liquid detergent. The SSL uses Liqui Nox.
• Dow Saran F-310 Resin, available from Dow Chemical Company.
• Plastic lacquer. Prepare plastic lacquer with resin to solvent ratios of 1:4 and 1:7 on a
weight basis. Fill a 3.8-L (1-gal) metal paint can with 2700 ±200 mL of solvent. Fill to the
bottom of handle rivet. Add 540 or 305 g of resin to make 1:4 or 1:7 plastic lacquer,
respectively. For the initial field and laboratory coatings, use the 1:4 plastic lacquer. Use
1:7 plastic lacquer for the last two laboratory coats. The 1:7 plastic lacquer is used to
conserve the resin and to reduce cost. In the field, mix solvent with a wooden stick. In the
laboratory, stir solvent with a non-sparking, high-speed stirrer while slowly adding resin.
Stir plastic lacquer for 15 min at 25°C. Store plastic lacquer in covered plastic or steel
containers. Acetone may be substituted for MEK.
5.24.6 Procedure
1. Prepare a round stock tag with sample identification number. Cut the copper wire and loop
around the clod. Record weight (TAG) of tag and wire. Loop fine copper wire around clod,
leaving a tail to which round stock tag is attached. Record weight of clod (CC1).
2. Dip clod in 1:4 plastic lacquer. Wait 7 min and then dip clod in 1:7 plastic lacquer. Wait 12
min and then dip clod in 1:7 plastic lacquer. Wait 55 min and then reweigh clod. If clod has
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adsorbed >3% plastic by clod weight or smells excessively of solvent, allow longer drying
time, then reweigh clod and record weight (CC2).
3. Dry clod in an oven at 110°C until weight is constant. Weigh oven-dry clod in air (WODC)
and in water (WODCW) and record weights.
4. If clod contains >5% rock fragments by weight, remove them from clod. Place clod in a
beaker and place on hot plate. Cover hot plate with a liquid vapor trap. Use a fume hood.
Heat clod on hot plate in excess of 200°C for 3 to 4 h. The plastic coating disintegrates at
temperatures above 200°C. After heating, clod should appear black and charred. Remove
clod from hot plate, lightly coat with liquid detergent, and add hot water.
5. Wet sieve the cool soil through a 2-mm, square- hole sieve. Dry and record weight (RF) of
rock fragments that are retained on the sieve. Determine rock fragment density by weighing
them in air to obtain their mass and in water to obtain their volume. If rock fragments are
porous and have a density similar to soil sample, do not correct clod mass and volume
measurement for rock fragments. If rock fragments are porous and have a density similar to
soil sample, do not correct clod mass and volume measurement for rock fragments. Correct
for rock fragments if these fragments can withstand breakdown when dry soil is placed
abruptly in water or calgon.
6. Correct bulk density for weight and volume of plastic coating. The coating has an air-dry
density of« 1.3 g cm-3. The coating loses 10 to 20% of its air-dry weight when dried in oven
at110°C.
5.24.7 Calculations
Dbod = [WODC- RF-ODPC - TAG]/{[(WODC - WODCW)/WD] - (RF/PD) - (MPC/1.3)}
where:
Dbod = Bulk density in g cm-3 of <2 mm, oven-dry fabric
WODC = Weight of oven-dry coated clod
RF = Weight of rock fragments
TAG = Weight of tag and wire
ODPC = MPC1 x 0.85, weight of oven-dry plastic coat
WD = Water density
PD = Density of rock fragments
WODCW = Weight of oven-dry coated clod in water
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MPC = [(CC2 - CC1) + FCE] x RV
where:
MPC = Weight of plastic coat before oven-drying
CC2 = Weight of clod after three laboratory plastic coats
CC1 = Weight of clod before three laboratory plastic coats
RV = Percent estimate of remaining clod volume after cutting to obtain flat surface
(« 80%)
FCE= 1.5x[(CC2-CC1)/3]
where:
FCE = Estimate of field-applied plastic coat
5.24.8 Report
Bulk density is reported to the nearest 0.01g cm"3.
5.24.9 References
Blake, G.R., and K.H. Hartge. 1986. Bulk density, p. 363-382. In A. Klute (ed.) Methods of soil
analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9. ASA
andSSSA, Madison, Wl.
Brasher, B.R., D.P. Franzmeier, V.T. Volassis, and S.E. Davidson. 1966. Use of saran resin to
coat natural soil clods for bulk density and water retention measurements. Soil Sci.
101:108.
Grossman, R.B., andT.G. Reinsch. 2002. Bulk density and linear extensibility, p. 201-228. In
J.H. Dane and G.C. Topp (eds.) Methods of soil analysis, Part 4. Physical methods. Soil
Sci. Am. Book Series No. 5. ASA and SSSA, Madison, Wl.
Occupational Safety and Health Administration. 1989. Air contaminants; final rule. Federal
Register. 29 CFR Part 1910. Vol. 54. No. 12. p. 2452-2453. U.S. Govt. Print. Office,
Washington, DC.
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5.25 Bulk Density (33-kPA Desorption, clods only)
5.25.1 Summary of Method
Field-occurring fabric (clods only) are collected from the face of an excavation. One coat of
plastic lacquer is applied in the field. Additional coats of plastic lacquer are applied in the
laboratory. The clod is desorbed to 33 kPa. After equilibration, the clod is weighed in air to
measure its mass and in water to measure its volume. After the clod is dried in the oven at
110°C, its mass and volume are determined again. A correction is made for the mass and
volume of rock fragments and for plastic coatings (Brasher et al., 1966; Blake and Hartge,
1986).
5.25.2 Interferences
Errors are caused by non-representative samples. Only field-occurring fabric (clods) should be
sampled. The whole soil bulk density may be overestimated because sampled clods frequently
exclude the crack space between clods (Grossman and Reinsch, 2002).
The penetration of plastic lacquer into the voids of sandy and organic soils interferes with the
corrections for mass and volume of the plastic coat and with the accuracy of water content
determinations. Penetration can be reduced by spraying water on the clod followed by
immediately dipping the clod in the plastic lacquer. Dipping should be done as quickly as
possible to reduce penetration of plastic.
Loss of soil during the procedure will void the analyses because all calculations are based on
the oven-dry soil mass. Holes in the plastic coating, which are detected by escaping air bubbles
from submerged clod, introduce errors in volume measurement. An inadequate evaporation of
plastic solvent results in overestimation of the soil mass. A drying time of 1 h is usually
sufficient time for evaporation of solvent. However, clods with high organic matter content may
need to dry longer.
Clods placed in an unsealed plastic bag can lose moisture during storage prior to analysis. If
clods irreversibly dry below 33-kPa-water content, then Db1/3 values for 33 kPa will be
erroneous. Completely seal the plastic storage bag to prevent drying.
Bulk density is reported for <2-mm soil fabric. Correction for rock fragments >2-mm requires
either knowledge or assumption of the rock fragment density. Estimate or measurement errors
of rock fragment density will affect the accuracy of the value for soil bulk density. Rock
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fragments may contain water, which complicates the application to actual water-holding
capacity.
5.25.3 Safety
Methyl ethyl ketone (MEK) is extremely flammable. A type B fire extinguisher should be in close
proximity in the laboratory. No open flames or nearby operation of electrical equipment are
permitted while using MEK. The MEK vapor is classified as a sensory irritant. The 8-h time
weighted average (TWA) exposure limit is 200 ppm, and the short-term exposure limit (STEL) is
300 ppm (Occupational Safety and Health Administration, 1989). Avoid physical contact. Use
with adequate ventilation. In closed buildings, use a fume hood. Keep in tightly closed
containers. Use safety glasses, proper gloves, and a lab coat. Wash hands immediately after
handling MEK. Additional information on the safe handling of MEK is available in Chemical
Safety Data Sheet SD-83, Manufacturing Chemists' Association, Inc., 1825 Connecticut Ave.
NW, Washington, DC.
Saran F-310 resin will decompose rapidly at temperatures >200°C releasing hydrogen chloride
gas. Avoid contact with Fe, Zn, Cu, and Al in solution. Avoid all contact with strong bases.
5.25.4 Equipment
• Electronic balance, ±0.01-g sensitivity
• Pressure plate extractor with porous ceramic plate.
• Air pressure, 33-kPa
• Rigid shipping containers. The SSL uses a corrugated box with compartments.
• Plastic bags, 1 mil, 127 x 89 x 330 mm
• Wire. The SSL uses a 28-awg coated copper wire.
• Hairnets
• Stock tags, 25.4-mm (1-in) diameter paper tag, with metal rim
• Hook assembly for weighing below balance
• Plexiglass water tank mounted on a fulcrum and lever to elevate tank
Oven, 110°C
• Sieve, No. 10 (2 mm openings)
• Rope, 3 m
• Clothespins
Knife
• Tile cut off saw with diamond blade
• Silt loam soil
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• Hot plate
• Desiccator with ceramic plate
• Vacuum, 80 kPa (0.8 bar)
• Metal probe
• Spray bottle
• Liquid vapor trap. The SSL constructs a tin enclosure over hot plate with a chimney and
duct to transfer vapor to water stream.
• Reinforced paper towels or cheesecloth
• Tension table. The SSL constructs a tension table by placing porous firebricks, covered
with reinforced paper towels, in a tub of water.
5.25.5 Reagents
• Methyl ethyl ketone (MEK), practical (2-butanone)
• Water
• Alcohol
• Liquid detergent. The SSL uses Liqui Nox.
• Dow Saran F-310 Resin, available from Dow Chemical Company.
• Plastic lacquer. Prepare plastic lacquer with resin to solvent ratios of 1:4 and 1:7 on a
weight basis. Fill a 3.8-L (1-gal) metal paint can with 2700 ±200 mL of solvent. Fill to the
bottom of handle rivet. Add 540 or 305 g of resin to make 1:4 or 1:7 plastic lacquer,
respectively. For the initial field and laboratory coatings, use the 1:4 plastic lacquer. Use
1:7 plastic lacquer for the last two laboratory coats. The 1:7 plastic lacquer is used to
conserve the resin and to reduce cost. In the field, mix solvent with a wooden stick. In the
laboratory, stir solvent with a non-sparking, high-speed stirrer while slowly adding resin.
Stir plastic lacquer for 15 min at 25°C. Store plastic lacquer in covered plastic or steel
containers. Acetone may be substituted for MEK.
5.25.6 Procedure
1. Prepare a round stock tag with sample identification number. Cut the copper wire and loop
around the clod (Fig. 1). Record weight (TAG) of tag and wire. Loop fine copper wire
around clod, leaving a tail to which round stock tag is attached. Record weight of clod
(CC1).
2. Dip clod in 1:4 plastic lacquer. Wait 7 min and then dip clod in 1:7 plastic lacquer. Wait 12
min and then dip clod in 1:7 plastic lacquer. Wait 55 min and then reweigh clod. If clod has
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adsorbed >3% plastic by clod weight or smells excessively of solvent, allow longer drying
time, then reweigh clod and record weight (CC2).
3. With a diamond saw, cut a flat surface on the clod. Place cut clod surface on a tension
table, maintained at 5 cm tension (Fig.2). Periodically check clod to determine if it has
reached equilibrium by inserting metal probe, touching, or by weight comparison. When
clod has reached equilibrium, remove clod and record weight (WSC).
4. If cut clod does not adsorb water, place clod in a desiccator on a water-covered plate with a
0-cm tension. Submerge only the surface of clod in the water. Add a few ml_ of alcohol.
Use in-house vacuum and apply suction until clod has equilibrated at saturation. Remove
clod and record weight (WSC).
5. Place clod in a pressure plate extractor. To provide good contact between clod and ceramic
plate, cover ceramic plate with a 5 mm layer of silt loam soil and saturate with water. Place
a sheet of reinforced paper towel or cheesecloth over the silt loam soil. Place surface of cut
clod on paper towel. Close container and secure lid. Apply gauged air pressure of 33 kPa.
When water ceases to discharge from outflow tube, clod is at equilibrium. Extraction usually
takes 3 to 4 wk. Remove clod and record weight (WMC). Compare WMC to WSC. If WMC
> WSC, equilibrate clod on tension table and repeat desorption process.
6. Dip clod in the 1:4 plastic lacquer. Wait 7 min and then dip clod in 1:7 plastic lacquer. Wait
12 min and then dip clod in 1:7 plastic lacquer. Wait 12 min and dip clod in 1:7 plastic
lacquer. After 55 min, reweigh clod and record weight (CCS). If clod has adsorbed >3%
plastic by weight or smells excessively of solvent, allow longer drying time, then reweigh
clod.
7. The clod should be waterproof and ready for volume measurement by water displacement.
Suspend clod below the balance, submerge in water, and record weight (WMCW).
8. Dry clod in an oven at 110°C until weight is constant. Weigh oven-dry clod in air (WODC)
and in water (WODCW) and record weights.
9. If clod contains >5% rock fragments by weight, remove them from clod. Place clod in a
beaker and place on hot plate. Cover hot plate with a liquid vapor trap. Use a fume hood.
Heat clod on hot plate in excess of 200°C for 3 to 4 h. The plastic coating disintegrates at
temperatures above 200°C. After heating, clod should appear black and charred. Remove
clod from hot plate, lightly coat with liquid detergent, and add hot water.
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10. Wet sieve the cool soil through a 2-mm, square-hole sieve. Dry and record weight (RF) of
rock fragments that are retained on the sieve. Determine rock fragment density by weighing
them in air to obtain their mass and in water to obtain their volume. If rock fragments are
porous and have a density similar to soil sample, do not correct clod mass and volume
measurement for rock fragments. Correct for rock fragments if these fragments can
withstand breakdown when dry soil is placed abruptly in water or calgon.
11. Correct bulk density for weight and volume of plastic coating. The coating has an air-dry
density of ~ 1.3g cm"3. The coating loses 10 to 20% of its air-dry weight when dried in oven
at110°C.
5.25.7 Calculations
Db33 = [WODC - RF - ODPC - TAG]/{[(CC3 - WMCW)/WD] - (RF/PD) - (MPC1/1.3)}
where:
Db33 = Bulk density in g cc-1 of <2 mm fabric at 33 kPa tension
WODC= Weight of oven-dry coated clod
RF = Weight of rock fragments
TAG = Weight of tag and wire
ODPC = MPC1 x 0.85, weight of oven-dry plastic coat
CCS = Weight of equilibrated clod after four additional plastic coats
WD = Water density
PD = Density of rock fragments
MPC1 = Weight of plastic coat before oven-drying
WMCW = Weight in water of coated clod equilibrated at 33-kPa tension
MPC1 = {[(CC2 - CC1) + FCE] x RV} + (CC3-WMC)
where:
MPC1 = Weight of plastic coat before oven-drying
CC2 = Weight of clod after three laboratory plastic coats
CC1 = Weight of clod before three laboratory plastic coats
WMC = Weight of coated clod equilibrated at 33-kPa tension
RV = Percent estimate of remaining clod volume after cutting to obtain flat
surface (« 80%)
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FCE= 1.5x[(CC2-CC1)/3]
where:
FCE = Estimate of field-applied plastic coat
Dbod = [WODC- RF- ODPC - TAG]/ {[(WODC - WODCW)/WD] - (RF/PD) - (MPC 1/1.3)}
where:
Dbod = Bulk density in g cc-1 <2 mm fabric, oven-dry fabric
WODCW = Weight of oven-dry clod coated in water
W33 ={[(CC3 - MPC1) - (WODC - ODPC)]/[(WODC - RF - ODPC - TAG)]} x 100
where:
W33 = Percent water weight retained at 33-kPa tension
5.25.8 Report
Bulk density is reported to the nearest 0.01g cm"3.
5.25.9 References
Blake, G.R., and K.H. Hartge. 1986. Bulk density, p. 363-382. In A. Klute (ed.) Methods of soil
analysis. Part 1. Physical and mineralogical methods. 2nd ed. Agron. Monogr. 9. ASA
andSSSA, Madison, Wl.
Brasher, B.R., D.P. Franzmeier, V.T. Volassis, and S.E. Davidson. 1966. Use of saran resin to
coat natural soil clods for bulk density and water retention measurements. Soil Sci.
101:108.
Grossman, R.B. and T.G. Reinsch. 2002. Bulk density and linear extensibility, p. 201-228. In
J.H. Dane and G.C. Topp (eds.) Methods of soil analysis, Part 4. Physical methods. Soil
Sci. Am. Book Series No. 5. ASA and SSSA, Madison, Wl.
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Occupational Safety and Health Administration. 1989. Air contaminants; final rule. Federal
Register. 29 CFR Part 1910. Vol. 54. No. 12. p. 2452-2453. U.S. Govt. Print. Office,
Washington, DC.
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Appendix A
Sample Laboratory Forms
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Page 196 of 256
Example Tracking Form
Date
Received
Sample
Type
Sample ID
Number
Site ID
Number
Laboratory
Tracking #
Notes
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Water Chemistry Bench Sheet
Project Name/Number,
Analyst_
11
o
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Water Chemistry Quality Assurance Bench Sheet
Project Name/Number,
Analyst_
BATCH ID
SAMPLEID
REPNUMBER
MATRIX
RESULT TYPE
PARAMETER
MEASURE
UNIT
DILUTION FACTOR
TRUEVALUE
PERCENT
PERCENTTYPE
ACCEPTANCERANGE
FLAG
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Diatom Enumeration Data Sheet
Site ID #
Lab#
Sample ID#
Date Collected
Slide Prepared by.
Concentration Factor
Volume
Steps,
Slide Identified by.
Notes
Taxa
Count Data
Transect 1
Transect 2
Transect 3
Transect 4
Transect 5
Transect 6
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Soft Algae Enumeration Data Sheet
Site ID #
Date Collected
Taxa
Sample ID#
Prepared by
Natural Count
Count Data
Sedgewick-Rafter
Lab#
Identified by
Palmer- Maloney
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Chlorophyll a Data Sheet
Lab ID #
Analyst Initials
Comments
CHLA_DUP
CHLA
Corrected_CHLA_DU
Corrected_CHLA
Volume correction (m
Dilution factor
Raw_CHLA_DUP
Raw_CHLA
Flag
Filtered volume (ml)
Date analyzed
REGION
STATE/TRIBE
DATE_RECEIVED
DATE_COLLECTED
SAMPLE_TYPE
Lab ID
SAMPLEJD
VISIT_NO
SITE_ID
BATCH_ID
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National Wetlands Condition Assessment
Laboratory Methods Manual
Page 202 of 256
Chlorophyll a Quality Assurance Sheet
Lab ID #
Analyst Initials
BATCH ID
Rep Numbe
Lab ID
Sample Typt
Date analyze!
Filtered volume (ml)
Flacj
Extract Concentration
Extract units
Dilution facto
Comments
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Appendix B
NLA and NAWQA Algae Taxa Lists
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The NAWQA diatom and non-diatom taxa list can be found at the following website:
http://diatom.acnatsci.org/nawqa/taxalist.asp
NLA Taxa
NLA Diatom and Non-Diatom Taxa
Achnanthes arenaria Amosse
Achnanthes coarctata (Brebisson) Grunow
[Achnanthes convergens Kobayasi
Achnanthes curtissima Carter
Achnanthes delicatissima Simonsen
Achnanthes depressa (Cleve) Hustedt
[Achnanthes gracillima Hustedt
I
Achnanthes imperfecta Schimanski
Achnanthes impexa Lange-Bertalot
Achnanthes kriegeri Krasske
Achnanthes lemmermannii Hustedt
[Achnanthes levanderi Hustedt
I
Achnanthes minuscula Hustedt
Achnanthes nitidiformis Lange-Bertalot
Achnanthes nodosa Cleve
[Achnanthes nodosa Cleve
f-~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~^^
Achnanthes oblongella 0strup
Achnanthes pseudoswazi Carter
Achnanthes rosenstockii Lange-Bertalot
Achnanthes rossii Hustedt
[Achnanthes rostellata Cleve-Euler
I
Achnanthes spp.
Achnanthes stolida (Krasske) Krasske
Achnanthes subhudsonis var. kraeuselii (Cholnoky) Cholnoky
[Achnanthes submarina Hustedt
I
Achnanthes trinodis (Ralfs) Grunow
Achnanthes ziegleri Lange-Bertalot
Achnanthidium affine (Grunow) Czarnecki
[Achnanthidium altergracillima (Lange-Bertalot) Round et Bukhtiyarova
[Achnanthidium biasolettianum (Kutzing) Bukhtiyarova
Achnanthidium catenatum (Bily et Marvan) Lange-Bertalot
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Achnanthidium deflexum (Reimer) Kingston
I
[Achnanthidium eutrophilum (Lange-Bertalot) Lange-Bertalot
I
[Achnanthidium exiguum (Grunow) Czarnecki
jAchnanthidium exiguum van. 1 NLS PJG
JAchnanthidium exiguum van. constrictum (Grunow) Andresen, Stoermer et Kreis
Achnanthidium exiguum van. elliptica Hustedt
I
[Achnanthidium exiguum van. heterovalvum (Krasske) Czarnecki
jAchnanthidium exilis (Kutzing) Round et Bukhtiyarova
JAchnanthidium lapidosum (Krasske) Kobayasi
Achnanthidium macrocephalum (Hustedt) Round et Bukhtiyarova
I ~
[Achnanthidium minutissimum (Kutzing) Czarnecki
[Achnanthidium rivulare Potapova et Ponader
jAchnanthidium saprophila (Kobayasi et Mayama) Round et Bukhtiyarova
JAchnanthidium spp.
Achnanthidium strictum Reichardt
[Actinella punctata Lewis
jActinocyclus normanii (Gregory) Hustedt
JAdlafia bryophila (Petersen) Lange-Bertalot
Adlafia minuscula (Grunow) Lange-Bertalot
I
[Adlafia minuscula var. muralis (Grunow) Lange-Bertalot
JAdlafia multnomahii Morales et Le
JAdlafia spp.
JAdlafia suchlandtii (Hustedt) Lange-Bertalot
Amphicampa eruca Ehrenberg
I
[Amphipleura kriegeriana (Krasske) Hustedt
jAmphipleura pellucida (Kutzing) Kutzing
JAmphipleura spp.
Amphora acutiuscula Kutzing
I
[Amphora coffeaeformis (Agardh) Kutzing
[Amphora commutata Grunow
JAmphora copulata (Kutzing) Schoeman et Archibald
JAmphora delicatissima Krasske
Amphora exigua Gregory
.
[Amphora fogediana Krammer
[Amphora holsatica Hustedt
JAmphora montana Krasske
[Amphora ovalis (Kutzing) Kutzing
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Amphora pediculus (Kutzing) Grunow
[Amphora robusta Gregory
[Amphora spp.
JAmphora strigosa Hustedt
JAmphora subcapitata (Kisselev) Hustedt
Amphora thumensis (Mayer) Krieger
.
[Amphora veneta Kutzing
JAneumastus minor (Hustedt) Lange-Bertalot
JAneumastus tusculus (Ehrenberg) Mann et Stickle
Anomoeoneis costata (Kutzing) Hustedt
I
[Anomoeoneis follis (Ehrenberg) Cleve
I
[Anomoeoneis sphaerophora (Kutzing) Pfitzer
JAnomoeoneis sphaerophora fo. costata (Kutzing) Schmidt
JAnomoeoneis sphaerophora fo. sculpta (Ehrenberg) Krammer
Anomoeoneis spp.
I
[Anomoeoneis styriaca (Grunow) Hustedt
JAnorthoneis excentrica (Donkin) Grunow
JAstartiella bahusiensis (Grunow) Witkowski, Lange-Bertalot et Metzeltin
Asterionella formosa Hassal
[Asterionella ralfsii Smith
jAsterionella spp.
JAulacoseira alpigena (Grunow) Krammer
JAulacoseira ambigua (Grunow) Simonsen
Aulacoseira canadensis (Hustedt) Simonsen
[Aulacoseira crassipunctata Krammer
JAulacoseira crenulata (Ehrenberg) Thwaites
JAulacoseira distans var. nivalis (Smith) Haworth
Aulacoseira distans var. nivaloides (Camburn) Haworth
I
[Aulacoseira granulata (Ehrenberg) Simonsen
jAulacoseira granulata fo. valida (Hustedt) Simonsen
JAulacoseira granulata var. angustissima (Miiller) Simonsen
JAulacoseira herzogii (Lemmermann) Simonsen
Aulacoseira islandica (Miiller) Simonsen
I ~
[Aulacoseira italica var. tenuissima (Grunow) Simonsen
JAulacoseira italica var. valida (Grunow) Simonsen
JAulacoseira lacustris (Grunow) Krammer
JAulacoseira laevissima (Grunow) Krammer
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Aulacoseira lirata (Ehrenberg) Ross
[Aulacoseira nygaardii Camburn
f-~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~~^^
Aulacoseira perglabra (0strup) Haworth
JAulacoseira perglabra var. florinae (Camburn) Haworth
JAulacoseira pfaffiana (Reinsch) Krammer
Aulacoseira spp.
I -
[Aulacoseira subarctica (Muller) Haworth
JAulacoseira tenella (Nygaard) Simonsen
JAulacoseira tenuior (Grunow) Krammer
Aulacoseira tethera Haworth
I
[Aulacoseira valida (Grunow) Krammer
Bacillaria paradoxa Gmelin
Bacillaria spp.
Biremis circumtexta (Meister ex Hustedt) Lange-Bertalot et Witkowski
Brachysira apiculata (Boyer) Lange-Bertalot et Moser
Brachysira aponina Kutzing
Brachysira brebissonii Ross
Brachysira elliptica Metzeltin et Lange-Bertalot
Brachysira microcephala (Grunow) Compere
Brachysira serians (Brebisson) Round et Mann
Brachysira spp.
Caloneis alpestris (Grunow) Cleve
Caloneis amphisbaena (Bory) Cleve
Caloneis bacillum (Grunow) Cleve
Caloneis borealis Carter
Caloneis branderii (Hustedt) Krammer
Caloneis clevei (Lagerstedt) Cleve
Caloneis hyalina Hustedt
Caloneis lewisii Patrick
Caloneis macedonica (Gregory) Krammer
Caloneis molaris (Grunow) Krammer
Caloneis schumanniana (Grunow) Cleve
Caloneis schumanniana var. biconstricta (Grunow) Reichelt
Caloneis silicula (Ehrenberg) Cleve
Caloneis spp.
Caloneis tenuis (Gregory) Krammer
Caloneis undulata (Gregory) Krammer
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Caloneis ventricosa van. subundulata (Grunow) Patrick
Caloneis westii (Smith) Hendey
Campylodiscus bicostatus Smith
Campylodiscus clypeus Ehrenberg
Campylodiscus hibernicus Ehrenberg
Campylodiscus levanderi Hustedt
Campylodiscus noricus Ehrenberg
Campylodiscus spp.
Campylopyxis germainii Mann
Campylostylus normannianus (Greville) Gerloff, Natour et Rivera
Capartogramma crucicula (Grunow ex Cleve) Ross
Cavinula cocconeiformis (Gregory ex Greville) Mann et Stickle
Cavinula jaernefelti (Hustedt) Mann et Stickle
Cavinula lacustris (Gregory) Mann et Stickle
Cavinula lapidosa (Krasske) Lange-Bertalot
Cavinula pseudoscutiformis (Grunow ex Schmidt) Mann et Stickle
Cavinula pusio (Cleve) Lange-Bertalot
Cavinula scutelloides (Smith) Lange-Bertalot et Metzeltin
Cavinula scutiformis (Grunow ex Schmidt) Mann et Stickle
Cavinula spp.
Cavinula variostriata (Krasske) Lange-Bertalot
Cavinula weinzierlii (Schimanski) Czarnecki
Chaetoceros spp.
Chamaepinnularia begeri (Krasske) Lange-Bertalot
Chamaepinnularia bremensis (Hustedt) Lange-Bertalot
Chamaepinnularia evanida (Hustedt) Lange-Bertalot
Chamaepinnularia krookii (Grunow) Lange-Bertalot et Krammer
Chamaepinnularia mediocris (Krasske) Lange-Bertalot
Chamaepinnularia schauppiana Lange-Bertalot et Metzeltin
Chamaepinnularia soehrensis (Krasske) Lange-Bertalot et Krammer
Chamaepinnularia soehrensis var. hassiaca (Krasske) Lange-Bertalot
Chamaepinnularia soehrensis var. muscicola (Petersen) Lange-Bertalot et Krammer
Chamaepinnularia sp. 1 NLS GDL
Chamaepinnularia spp.
Cocconeis diminuta Pantocsek
Cocconeis disculus (Schumann) Cleve
Cocconeis neodiminuta Krammer
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Cocconeis neothumensis Krammer
Cocconeis pediculus Ehrenberg
Cocconeis peltoides Hustedt
Cocconeis placentula Ehrenberg
Cocconeis placentula var. klinoraphis Geitler
Cocconeis placentula var. rouxii (Heribaud et Brun in Heribaud) Cleve
Cocconeis pseudolineata (Geitler) Lange-Bertalot
Cocconeis pseudothumensis Reichardt
Cocconeis scutellum Ehrenberg
Cocconeis spp.
Coscinodiscus spp.
Cosmioneis spp.
Craticula accomoda (Hustedt) Mann
Craticula acidoclinata Lange-Bertalot et Metzeltin
Craticula ambigua (Ehrenberg) Mann
Craticula buderi (Hustedt) Lange-Bertalot
Craticula citrus (Krasske) Reichardt
Craticula cuspidata (Kutzing) Mann
Craticula halophila (Grunow) Mann
Craticula halophilioides (Hustedt) Lange-Bertalot
Craticula molestiformis (Hustedt) Lange-Bertalot
Craticula riparia (Hustedt) Lange-Bertalot
Craticula riparia var. mollenhaueri Lange-Bertalot
Craticula sardiniensis (Lange-Bertalot, Cavacini, Tagliaventi et Alfinito) Morales et Le
Craticula spp.
Craticula submolesta (Hustedt) Lange-Bertalot
Craticula vixnegligenda Lange-Bertalot
Ctenophora pulchella (Ralfs ex Kutzing) Williams et Round
Ctenophora pulchella var. lacerata (Hustedt) Bukhtiyarova
Cyclostephanos costatilimbus Stoermer, Hakansson etTheriot
Cyclostephanos damasii (Hustedt) Stoermer et Hakansson
Cyclostephanos dubius (Fricke) Round
Cyclostephanos invisitatus (Hohn et Hellerman) Theriot, Stoermer et Hakansson
Cyclostephanos spp.
Cyclostephanos tholiformis Stoermer, Hakansson et Theriot
Cyclotella antiqua Smith
Cyclotella atomus Hustedt
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Cyclotella bodanica van. affinis (Grunow) Cleve-Euler
Cyclotella bodanica van. lemanica O. Mullen (Bachmann)
Cyclotella comensis Grunow et Van Heurck
Cyclotella cyclopuncta Hakansson
Cyclotella delicatula Hustedt
Cyclotella distinguenda Hustedt
Cyclotella distinguenda van. unipunctata (Hustedt) Hakansson et Carter
Cyclotella gamma Skvortzow
Cyclotella glabriuscula (Grunow) Hakansson
Cyclotella hakanssoniae Wendker
Cyclotella krammeri Hakansson
Cyclotella kuetzingiana Thwaites
Cyclotella meneghiniana Kutzing
Cyclotella michiganiana Skvortzow
Cyclotella michiganiana var. 1 NLS GDI
Cyclotella ocellata Pantocsek
Cyclotella planctonica Brunnthaler
Cyclotella polymorpha Meyer et Hakansson
Cyclotella quadrijuncta (Schroter) von Keissler
Cyclotella schumannii (Grunow) Hakansson
Cyclotella spp.
Cyclotella striata (Kutzing) Grunow
Cyclotella tripartita Hakansson
Cylindrotheca gracilis (Brebisson) Grunow
Cymatopleura elliptica (Brebisson) Smith
Cymatopleura solea (Brebisson) Smith
Cymatopleura spp.
Cymbella acuta (Schmidt in Schmidt et al.) Cleve
Cymbella aequalis Smith
Cymbella affinis Kutzing
Cymbella alpina Grunow
Cymbella amphicephala Na'geli ex Kutzing
Cymbella amphioxys (Kutzing) Cleve
Cymbella angustata (Smith) Cleve
Cymbella aspera (Ehrenberg) Peragallo
Cymbella austriaca Grunow
Cymbella budayana Pantocsek
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Cymbella caespitosa Brun
Cymbella cistula (Ehrenberg) Kirchner
Cymbella cuspidata Kutzing
Cymbella cymbiformis Agardh
Cymbella delicatula Kutzing
Cymbella diluviana (Krasske) Florin
Cymbella ehrenbergii Kutzing
Cymbella elginensis Krammer
Cymbella gracilis (Ehrenberg) Kutzing
Cymbella hauckii Van Heurck
Cymbella helvetica Kutzing
Cymbella heteropleura (Ehrenberg) Kutzing
Cymbella hustedtii Krasske
Cymbella hybrida Grunow ex Cleve
Cymbella incerta (Grunow) Cleve
Cymbella incerta var. crassipunctata Krammer
Cymbella laevis Na'geli ex Kutzing
Cymbella lanceolata (Agardh) Agardh
Cymbella lapponica Grunow
Cymbella lata Grunow
Cymbella leptoceros (Ehrenberg) Kutzing
Cymbella mesiana Cholnoky
Cymbella mexicana (Ehrenberg) Cleve
Cymbella mexicana var. janischii (Schmidt) Reimer
Cymbella moelleriana Grunow
Cymbella naviculiformis Auerswald ex Heribaud
Cymbella obscura Krasske
Cymbella paucistriata Cleve-Euler
Cymbella proxima Reimer
Cymbella pusilla Grunow
Cymbella reinhardtii Grunow
Cymbella rupicola Grunow
Cymbella similis Krasske
Cymbella simonsenii Krammer
Cymbella smithii Rabenhorst
Cymbella spp.
Cymbella stauroneiformis Lagerstedt
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Cymbella subaequalisfo. krasskei (Foged) Reimer
Cymbella subaequalis Grunow
Cymbella subcuspidata Krammer
Cymbella subturgidula Krammer
Cymbella tumidula Grunow ex Schmidt
Cymbella tumidula var. lancettula Krammer
Cymbella turgida Gregory
Cymbella turgidula Grunow
Cymbella tynnii Krammer
Cymbellonitzschia diluviana Hustedt
Cymbopleura cuspidata Kutzing
Decussata placenta (Ehrenberg) Lange-Bertalot et Metzeltin
Denticula elegans Kutzing
Denticula kuetzingii Grunow
Denticula kuetzingii Grunow
Denticula spp.
Denticula subtilis Grunow
Denticula tenuis Kutzing
Denticula valida (Pedicino) Grunow
Diadesmis confervacea Kutzing
Diadesmis contenta (Grunow ex Van Heurck) Mann
Diadesmis contenta var. biceps (Grunow) Hamilton
Diadesmis gallica Smith
Diadesmis irata (Krasske) Moser, Lange-Bertalot et Metzeltin
Diadesmis laevissima (Cleve) Mann
Diadesmis perpusilla (Grunow) Mann
Diadesmis spp.
Diatoma anceps (Ehrenberg) Kirchner
Diatoma hyemalis (Roth) Heiberg
Diatoma mesodon (Ehrenberg) Kutzing
Diatoma moniliformis Kutzing
Diatoma spp.
Diatoma tenuis Agardh
Diatoma tenuis var. elongatum Lyngbye
Diatoma vulgaris Bory
Diatoma vulgaris var. breve Grunow
Diploneis boldtiana Cleve
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Diploneis elliptica (Kutzing) Cleve
Diploneisfinnica (Ehrenberg) Cleve
Diploneis litoralis (Donkin) Cleve
Diploneis marginestriata Hustedt
Diploneis modica Hustedt
Diploneis oblongella (Na'geli ex Kutzing) Ross
Diploneis oculata (Brebisson) Cleve
Diploneis ovalis (Hilse ex Rabenhorst) Cleve
Diploneis parma Cleve
Diploneis parma Cleve
Diploneis peterseni Hustedt
Diploneis pseudovalis Hustedt
Diploneis puella (Schumann) Cleve
Diploneis smithii (Brebisson ex Smith) Cleve
Diploneis smithii var. pumila (Grunow) Hustedt
Diploneis spp.
Diploneis subovalis Cleve
Discostella asterocostata (Xie, Lin et Cai) Houk et Klee
Discostella glomerata (Bachmann) Houk et Klee
Discostella spp.
Discostella stelligera (Hustedt) Houk et Klee
Ellerbeckia arenaria (Moore) Crawford
Encyonema auerswaldii Rabenhorst
Encyonema gaeumanii (Meister) Krammer
Encyonema gracile Rabenhorst
Encyonema hebridicum Grunow ex Cleve
Encyonema latens (Krasske) Mann
Encyonema lunatum (Smith) Van Heurck
Encyonema minutum (Hilse) Mann
Encyonema muelleri (Hustedt) Mann
Encyonema norvegicum (Grunow) Mills
Encyonema norvegicum (Grunow) Mills
Encyonema obscurum (Krasske) Mann
Encyonema obscurum var. alpina Krammer
Encyonema paucistriatum (Cleve-Euler) Mann
Encyonema perpusillum (Cleve) Mann
Encyonema prostratum (Berkeley) Kutzing
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Encyonema reichardtii (Krammer) Mann
Encyonema spp.
Encyonema triangulum (Ehrenberg) Kutzing
Encyonema tumida (Brebisson ex Kutzing) Mann
Encyonema ventricosum Agardh
Encyonopsis cesatii (Rabenhorst) Krammer
Encyonopsis delicatissima (Hustedt) Krammer
Encyonopsis descripta (Hustedt) Krammer
Encyonopsis evergladianum Krammer
Encyonopsis falaisensis (Grunow) Krammer
Encyonopsis krammeri Reichardt
Encyonopsis microcephala (Grunow) Krammer
Encyonopsis minuta Krammer et Reichardt
Encyonopsis spp.
Encyonopsis subminuta Krammer et Reichardt
Entomoneis alata (Ehrenberg) Ehrenberg
Entomoneis costata (Hustedt) Reimer
Entomoneis ornata (Bailey) Reimer
Entomoneis spp.
Epithemia adnata (Kutzing) Brebisson
Epithemia adnata var. porcellus (Kutzing) Patrick
Epithemia argus (Ehrenberg) Kutzing
Epithemia reichelti Fricke
Epithemia smithii Carruthers
Epithemia sorex Kutzing
Epithemia spp.
Epithemia turgida (Ehrenberg) Kutzing
Epithemia turgida var. granulata (Ehrenberg) Hustedt
Epithemia turgida var. westermannii (Ehrenberg) Grunow
Eucocconeis alpestris (Brun) Lange-Bertalot
Eucocconeisflexella (Kutzing) Cleve
Eucocconeis laevis (0strup) Lange-Bertalot
Eucocconeis laevis var. diluviana (Hustedt) Lange-Bertalot
Eucocconeis spp.
Eunotia arculus (Grunow ) Lange-Bertalot et Norpel
Eunotia arcus Ehrenberg
Eunotia bactriana Ehrenberg
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Eunotia bidentula Smith
Eunotia bigibba Kutzing
Eunotia bilunaris (Ehrenberg) Mills
Eunotia camelus Ehrenberg
Eunotia Carolina Patrick
Eunotia cf. paludosa NLA EWT Grunow
Eunotia cf. paludosa NLA JPK Grunow
Eunotia cf. pirla NLS DME Carter et Flower
Eunotia cf. sudetica NLA DB Mtiller
Eunotia circumborealis Lange-Bertalot et Norpel
Eunotia croatana Siver, Hamilton et Morales
Eunotia curvata fo. bergii Woodhead et Tweed
Eunotia diodon Ehrenberg
Eunotia elegans 0strup
Eunotia exigua (Brebisson ex Kutzing) Rabenhorst
Eunotia faba (Ehrenberg) Grunow
Eunotia fallax Cleve
Eunotia fallax var. gracillima Krasske
Eunotia fallax var. groenlandica (Grunow) Lange-Bertalot et Norpel
Eunotia flexuosa Brebisson ex Kutzing
Eunotia formica Ehrenberg
Eunotia glacialis Meister
Eunotia hemicyclus (Ehrenberg) Karsten
Eunotia hexaglyphis Ehrenberg
Eunotia implicata Norpel, Lange-Bertalot et Alles
Eunotia incisa Smith ex Gregory
Eunotia inflata (Grunow) Norpel-Schempp et Lange-Bertalot
Eunotia intermedia (Krasske ex Hustedt) Norpel et Lange-Bertalot
Eunotia lapponica Grunow ex Cleve
Eunotia luna Ehrenberg
Eunotia meisteri Hustedt
Eunotia microcephala Krasske ex Hustedt
Eunotia minor (Kutzing) Grunow
Eunotia muscicola var. tridentula Norpel et Lange-Bertalot
Eunotia naegelii Migula
Eunotia neofallax Norpel-Schempp et Lange-Bertalot
Eunotia nymanii Grunow
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Eunotia nymanniana Grunow (sensu Hustedt)
Eunotia paludosa Grunow
Eunotia paludosa van. trinacria (Krasske) Norpel
Eunotia parallela Ehrenberg
Eunotia pectinalis (Mullen) Rabenhorst
Eunotia perpusilla Grunow
Eunotia pirla Carter et Flower
Eunotia praerupta Ehrenberg
Eunotia praerupta var. bigibba (Kutzing) Grunow
Eunotia praerupta var. curta Grunow
Eunotia pseudoparalleloides (Grunow) Norpel-Schempp et Lange-Bertalot
Eunotia rabenhorstii Grunow
Eunotia rhomboidea Hustedt
Eunotia rhynchocephala Hustedt
Eunotia rhynchocephala var. satelles Norpel et Lange-Bertalot
Eunotia seminulum Norpel-Schempp et Lange-Bertalot
Eunotia septentrionalis 0strup
Eunotia serra Ehrenberg
Eunotia serra var. diadema (Ehrenberg) Patrick
Eunotia serra var. tetraodon Norpel
Eunotia silvahercynia Norpel, Van Sull et Lange-Bertalot
Eunotia siolii Hustedt
Eunotia soleirolii (Kutzing) Rabenhorst
Eunotia spp.
Eunotia subarcuatoides Alles, Norpel et Lange-Bertalot
Eunotia sudetica Muller
Eunotia tenella (Grunow) Hustedt
Eunotia triodon Ehrenberg
Eunotia valida Hustedt
Eunotia yanomami Metzeltin et Lange-Bertalot
Eunotia zasuminensis (Cabejszekowna) Korner
Eunotia zygodon Ehrenberg
Fallacia auriculata (Hustedt) Mann
Fallacia cryptolyra (Brockmann) Stickle et Mann
Fallacia enigmatica (Germain) Lange-Bertalot
Fallacia indifferens (Hustedt) Mann
Fallacia insociabilis (Krasske) Stickle et Mann
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Fallacia lenzii (Hustedt) Lange-Bertalot
Fallacia litoricola (Hustedt) Mann
Fallacia lucinensis (Hustedt) Mann in Round, Crawford et Mann
Fallacia monoculata (Hustedt) Mann
Fallacia omissa (Hustedt) Mann
Fallacia pygmaea (Kutzing) Stickle et Mann
Fallacia spp.
Fallacia subforcipata (Hustedt) Mann
Fallacia subhamulata (Grunow) Mann
Fallacia tenera (Hustedt) Mann
Fistulifera pelliculosa (Brebisson ex Kutzing) Lange-Bertalot
Fistulifera saprophila (Lange-Bertalot et Bonik) Lange-Bertalot
Fragilaria acidobiontica Charles
Fragilaria acutirostrata Metzeltin et Lange-Bertalot
Fragilaria alpestris Krasske
Fragilaria amphicephala (Kutzing) Lange-Bertalot
Fragilaria capucina Desmazieres
Fragilaria capucina var. mesolepta Rabenhorst
Fragilaria capucina var. perminuta (Grunow) Lange-Bertalot
Fragilaria construens var. exigua (Smith) Schulz
Fragilaria crotonensis Kitton
Fragilaria cyclopum (Brutschy) Lange-Bertalot
Fragilaria distans (Grunow) Bukhtiyarova
Fragilaria exigua Grunow
Fragilaria famelica (Kutzing) Lange-Bertalot
Fragilaria germainii Reichardt et Lange-Bertalot
Fragilaria heidenii 0strup
Fragilaria neoproducta Lange-Bertalot
Fragilaria nitzschioides Grunow
Fragilaria pinnata var. acuminata Mayer
Fragilaria radians (Kutzing) Lange-Bertalot
Fragilaria rhabdosoma Ehrenberg
Fragilaria sepes Ehrenberg
Fragilaria spp.
Fragilaria subconstricta 0strup
Fragilaria tenuistriata 0strup
Fragilaria vaucheriae (Kutzing) Petersen
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Fragilaria zeilleri van. elliptica Gasse
Fragilariforma bicapitata (Mayer) Williams et Round
Fragilariforma constricta (Ehrenberg) Williams et Round
Fragilariforma constricta fo. stricta (Cleve) Poulin
Fragilariforma constricta var. trinodis (Hustedt) Hamilton
Fragilariforma hungarica var. tumida (Cleve-Euler) Hamilton
Fragilariforma lata (Cleve-Euler) Williams et Round
Fragilariforma polygonata (Cleve-Euler) Kingston, Sherwood et Bengston
Fragilariforma spinulosa (Patrick) Metzeltin et Lange-Bertalot
Fragilariforma spp.
Fragilariforma virescens (Ralfs) Williams et Round
Frustulia amphipleuroides (Grunow) Cleve-Euler
Frustulia bahlsii Edlund et Brant
Frustulia crassinervia (Brebisson) Lange-Bertalot et Krammer
Frustulia erifuga Lange-Bertalot et Krammer
Frustulia krammeri Lange-Bertalot et Metzeltin
Frustulia pseudomagaliesmontana Camburn et Charles
Frustulia quadrisinuata Lange-Bertalot in Lange-Bertalot et Metzeltin
Frustulia saxonica Rabenhorst
Frustulia spp.
Frustulia viridula Kutzing
Frustulia vulgaris (Thwaites) DeToni
Frustulia weinholdii Hustedt
Geissleria acceptata (Hustedt) Lange-Bertalot et Metzeltin
Geissleria aikenensis (Patrick) Torgan et Olivera
Geissleria decussis (Hustedt) Lange-Bertalot et Metzeltin
Geissleria dolomitica (Bock) Lange-Bertalot et Metzeltin
Geissleria ignota (Krasske) Lange-Bertalot et Metzeltin
Geissleria paludosa (Hustedt) Lange-Bertalot et Metzeltin
Geissleria schoenfeldii (Hustedt) Lange-Bertalot et Metzeltin
Geissleria spp.
Gomphoneis eriense var. variabilis Kociolek et Stoermer
Gomphoneis geitleri Kociolek et Stoermer
Gomphoneis herculeana (Ehrenberg) Cleve
Gomphoneis herculeana var. robusta (Grunow) Cleve
Gomphoneis minuta Kociolek et Stoermer
Gomphoneis spp.
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Gomphoneis transsilvanica (Pantocsek) Krammer in Krammer et Lange-Bertalot
Gomphonema acuminatum Ehrenberg
Gomphonema acuminatum van. coronatum (Ehrenberg) Rabenhorst
Gomphonema acuminatum var. pusillum Grunow
Gomphonema acutiusculum (Mtiller) Cleve-Euler
Gomphonema affine Kutzing
Gomphonema amoenum Lange-Bertalot
Gomphonema angustatum (Kutzing) Rabenhorst
Gomphonema angustum Agardh
Gomphonema anjae Lange-Bertalot et Reichardt
Gomphonema apicatum Ehrenberg
Gomphonema apuncto Wallace
Gomphonema augur Ehrenberg
Gomphonema augur var. turris (Ehrenberg) Lange-Bertalot
Gomphonema auritum Braun
Gomphonema bohemicum Reichardt et Fricke
Gomphonema camburnii Lange-Bertalot
Gomphonema capitatum Ehrenberg
Gomphonema carolinense Hagelstein
Gomphonema clavatum Ehrenberg
Gomphonema clevei Fricke in Schmidt et al.
Gomphonema contraturris Lange-Bertalot et Reichardt
Gomphonema dichotomum Kutzing
Gomphonema drutelingense Reichardt
Gomphonema exilissimum (Grunow) Lange-Bertalot
Gomphonema germainii Kociolek et Stoermer
Gomphonema gibba Wallace
Gomphonema gracile Ehrenberg emend Van Heurck
Gomphonema gracile fo. cymbelloides (Grunow ms. in Cleve) Mayer
Gomphonema hebridense Gregory
Gomphonema innocens Reichardt
Gomphonema insigne Gregory
Gomphonema intricatum Kutzing
Gomphonema intricatum var. vibrio (Ehrenberg) Cleve
Gomphonema kobayashiae Metzeltin et Lange-Bertalot
Gomphonema kobayasii Kociolek et Kingston
Gomphonema lagenula Kutzing
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NLA Diatom and Non-Diatom Taxa
Gomphonema longilineare Reichardt
Gomphonema maclaughlinii Reichardt
Gomphonema mexicanum Grunow ex Van Heurck
Gomphonema micropumilum Reichardt
Gomphonema micropus Kutzing
Gomphonema minusculum Krasske
Gomphonema minutum (Agardh) Agardh
Gomphonema olivaceoides Hustedt
Gomphonema olivaceum (Lyngbye) Kutzing
Gomphonema parvulius (Lange-Bertalot et Reichardt) Lange-Bertalot et Reichardt
Gomphonema parvulum (Kutzing) Kutzing
Gomphonema patrickii Kociolek et Stoermer
Gomphonema patrickii Kociolek et Stoermer
Gomphonema productum (Grunow) Lange-Bertalot et Reichardt
Gomphonema pseudoaugur Lange-Bertalot
Gomphonema pseudotenellum Lange-Bertalot
Gomphonema pumilum (Grunow) Reichardt et Lange-Bertalot
Gomphonema pygmaeum Kociolek et Stoermer
Gomphonema quadripunctatum (0strup) Wislouch
Gomphonema rhombicum Fricke
Gomphonema sarcophagus Gregory
Gomphonema sphaerophorum Ehrenberg
Gomphonema spp.
Gomphonema subclavatum (Grunow) Grunow
Gomphonema subclavatum var. commutatum (Grunow) Mayer
Gomphonema subclavatum var. mexicanum (Grunow) Patrick
Gomphonema subtile Ehrenberg
Gomphonema subtile var. sagitta (Schumann) Grunow ex Van Heurck
Gomphonema truncatum Ehrenberg
Gomphonema truncatum var. capitatum (Ehrenberg) Patrick
Gomphonema turgidum Ehrenberg
Gomphonema turris Ehrenberg
Gomphonema utae Lange-Bertalot et Reichardt
Gomphonema variostriatum Camburn et Charles
Gomphonema vibrioides Reicherdt et Lange-Bertalot
Gomphonitzschia spp.
Gomphosphenia grovei (Schmid) Lange-Bertalot
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NLA Diatom and Non-Diatom Taxa
Gomphosphenia grovei van. lingulata (Hustedt) Lange-Bertalot
Gomphosphenia lingulatiformis (Lange-Bertalot et Reichardt) Lange-Bertalot
Gomphosphenia spp.
Gomphosphenia tackei (Hustedt) Lange-Bertalot
Gyrosigma acuminatum (Kutzing) Rabenhorst
Gyrosigma attenuatum (Kutzing) Rabenhorst
Gyrosigma balticum (Ehrenberg) Rabenhorst
Gyrosigma macrum (Smith) Griffith et Henfrey
Gyrosigma nodiferum (Grunow) Reimer
Gyrosigma obscurum (Smith) Griffith et Henfrey
Gyrosigma obtusatum (Sullivant et Wormley) Boyer
Gyrosigma parkerii (Harrison) Elmore
Gyrosigma scalproides (Rabenhorst) Cleve
Gyrosigma sciotoense (Sullivan et Wormley) Cleve
Gyrosigma spp.
Gyrosigma strigilis (Smith) Cleve
Hannaea arcus (Ehrenberg) Patrick
Hantzschia amphioxys (Ehrenberg) Grunow
Hantzschia distinctepunctata Hustedt
Hantzschia elongata (Hantzsch) Grunow
Hantzschia spectabilis (Ehrenberg) Hustedt
Hantzschia spp.
Hantzschia virgata (Roper) Grunow
Hantzschia vivax (Smith) Peragallo
Haslea spicula (Hickle) Lange-Bertalot
Hippodonta capitata (Ehrenberg) Lange-Bertalot, Metzeltin et Witkowski
Hippodonta costulata (Grunow) Lange-Bertalot, Metzeltin et Witkowski
Hippodonta hungarica (Grunow) Lange-Bertalot, Metzeltin et Witkowski
Hippodonta lueneburgensis (Grunow) Lange-Bertalot, Metzeltin et Witkowski
Hippodonta spp.
Hippodonta subcostulata (Hustedt) Lange-Bertalot, Metzeltin et Witkowski
Karayevia amoena (Hustedt) Bukhtiyarova
Karayevia bottnica (Cleve) Lange-Bertalot
Karayevia carissima (Lange-Bertalot) Bukhtiyarova
Karayevia clevei (Grunow) Bukhtiyarova
Karayevia clevei var. bottnica (Cleve) Bukhtiyarova
Karayevia clevei var. rostrata (Hustedt) Bukhtiyarova
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NLA Diatom and Non-Diatom Taxa
Karayevia laterostrata (Hantzsch) Bukhtiyarova
Karayevia ploenensis (Hustedt) Bukhtiyarova
Karayevia spp.
Karayevia suchlandtii (Hustedt) Bukhtiyarova
Kobayasia parasubtilissima (Kobayasi et Nagumo) Lange-Bertalot
Kobayasiella jaegii (Meister) Lange-Bertalot
Kobayasiella spp.
Kobayasiella subtilissima (Cleve) Lange-Bertalot
Kobayasiella venezuelensis Metzeltin et Lange-Bertalot
Krasskella kriegerana (Krasske) Ross et Sims
Lemnicola hungarica (Grunow) Round et Basson
Luticola cohnii (Hilse) Mann
Luticola dismutica (Hustedt) Mann in Round, Crawford et Mann
Luticola goeppertiana (Bleisch) Mann
Luticola mutica (Kutzing) Mann
Luticola muticoides (Hustedt) Mann
Luticola nivalis (Ehrenberg) Mann
Luticola nivaloides Johansen
Luticola spp.
Luticola ventricosa (Kutzing) Mann
Mastogloia elliptica (Agardh) Cleve
Mastogloia elliptica var. danseii (Thwaites) Cleve
Mastogloia grevillei Smith
Mastogloia lanceolata Thwaites in Smith
Mastogloia pumila (Cleve et Moller; Grunow in Van Heurck) Cleve
Mastogloia pusilla Grunow
Mastogloia smithii Thwaites
Mastogloia smithii var. lacustris Grunow
Mastogloia spp.
Mayamaea atomus (Kutzing) Lange-Bertalot
Mayamaea atomus var. permitis (Hustedt) Lange-Bertalot
Mayamaea excelsa (Krasske) Lange-Bertalot
Mayamaea recondita (Hustedt) Lange-Bertalot
Mayamaea spp.
Melosira arentii (Kolbe) Nagumo et Kobayasi
Melosira distans var. humilis Cleve-Euler
Melosira undulata (Ehrenberg) Kutzing
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NLA Diatom and Non-Diatom Taxa
Melosira varians Agardh
Meridion circulare (Greville) Agardh
Meridion circulare var. constrictum (Ralfs) Van Heurck
Meridion spp.
Microcostatus krasskei (Hustedt) Johansen et Sray
Microcostatus kuelbsii (Lange-Bertalot) Lange-Bertalot
Microcostatus maceria (Schimanski) Lange-Bertalot
Microcostatus naumannii (Hustedt) Lange-Bertalot
Muelleria gibbula (Cleve) Spaulding et Stoermer
Navicula aboensis (Cleve) Hustedt
Navicula absoluta Hustedt
Navicula amphibola Cleve
Navicula amphiceropsis Lange-Bertalot et Rumrich
Navicula angusta Grunow
Navicula angustata Smith
Navicula antonii Lange-Bertalot
Navicula arctotenelloides Lange-Bertalot et Metzeltin
Navicula arenaria Donkin
Navicula arvensis Hustedt
Navicula arvensis var. major Lange-Bertalot
Navicula aurora Sovereign
Navicula bacilloides Hustedt
Navicula bourrellyivera Lange-Bertalot, Witkowski et Stachura
Navicula brasiliana Cleve
Navicula canalis Patrick
Navicula capitatoradiata Germain
Navicula capitellata Skvortzow
Navicula cari Ehrenberg
Navicula catalanogermanica Lange-Bertalot et Hofmann
Navicula caterva Hohn et Hellerman
Navicula cincta (Ehrenberg) Ralfs
Navicula cocconeiformis var. capitata Krasske
Navicula comoides (Dillwyn; Dillwyn) Peragallo et Peragallo
Navicula concentrica Carter
Navicula constans Hustedt
Navicula constans var. symmetrica Hustedt
Navicula cryptocephala Kutzing
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NLA Diatom and Non-Diatom Taxa
Navicula cryptocephaloides Hustedt
Navicula cryptofallax Lange-Bertalot ex Hofmann
Navicula cryptotenella Lange-Bertalot in Krammer et Lange-Bertalot
Navicula denselineolata (Lange-Bertalot) Lange-Bertalot
Navicula detenta Hustedt
Navicula difficillima Hustedt
Navicula difficillimoides Hustedt
Navicula digitoradiata (Gregory) Ralfs ex Pritchard
Navicula digitoradiata var. rostrata Hustedt
Navicula digitulus Hustedt
Navicula disjuncta Hustedt
Navicula effrenata Krasske
Navicula eidrigiana Carter
Navicula erifuga Lange-Bertalot
Navicula exigua var. signata Hustedt
Navicula exilis Kutzing
Navicula facilis Krasske
Navicula farta Hustedt
Navicula festiva Krasske
Navicula fluens Hustedt
Navicula fossalis Krasske
Navicula fossalis var. obsidialis (Hustedt) Lange-Bertalot in Krammer et Lange-Bertalot
Navicula fracta Hustedt
Navicula gallica var. perpusilla (Grunow) Lange-Bertalot
Navicula gastrum var. signata Hustedt
Navicula gerloffi Schimanski
Navicula germainii Wallace
Navicula globulifera Hustedt
Navicula glomus Carter et Bailey-Watts
Navicula goersii Bahls
Navicula gottlandica Grunow
Navicula gregaria Donkin
Navicula hambergii Hustedt
Navicula hasta Pantocsek
Navicula heimansioides Lange-Bertalot
Navicula helensis Schulz
Navicula heufleriana (Grunow) Cleve
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NLA Diatom and Non-Diatom Taxa
Navicula hintzii Lange-Bertalot
Navicula hintzii Lange-Bertalot
Navicula hoeflerii Cholnoky
Navicula hustedtiana Simonsen
Navicula hustedtii van. obtusa Hustedt
Navicula ignota van. palustris (Hustedt) Lund
Navicula imbricata Bock
Navicula impexa Hustedt
Navicula incertata Hustedt
Navicula ingenua Hustedt
Navicula ingrata Krasske
Navicula integra (Smith) Ralfs
Navicula irmengardis Lange-Bertalot in Lange-Bertalot et Metzeltin
Navicula jaagii Meister
Navicula joubaudii Germain
Navicula kotschyi Grunow
Navicula kriegerii Krasske
Navicula kuelbsii Lange-Bertalot
Navicula lacustris Gregory
Navicula lanceolata (Agardh) Ehrenberg
Navicula lateropunctata Wallace
Navicula laterostrata Hustedt
Navicula laterostrata Hustedt
Navicula laticeps Hustedt
Navicula leistikowii Lange-Bertalot
Navicula leptostriata J0rgensen
Navicula lesmonensis Hustedt
Navicula levanderii Hustedt
Navicula libonensis Schoeman
Navicula lineolata Ehrenberg
Navicula longa (Gregory) Ralfs in Pritchard
Navicula longicephala Hustedt
Navicula ludloviana Schmidt in Schmidt et al.
Navicula lundii Reichardt
Navicula luzonensis Hustedt
Navicula mandumensis J0rgensen
Navicula margalithii Lange-Bertalot
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NLA Diatom and Non-Diatom Taxa
Navicula medioconvexa Hustedt
Navicula menisculus Schumann
Navicula menisculus van. obtusa Hustedt
Navicula meniscus Schumann
Navicula minima Grunow
Navicula minusculoides Hustedt
Navicula minuta Cleve-Euler
Navicula minutula Smith
Navicula modica Hustedt
Navicula moskalii Metzeltin, Witkowski et Lange-Bertalot
Navicula muraliformis Hustedt
Navicula normaloides Cholnoky
Navicula notha Wallace
Navicula oblonga 0strup
Navicula oblongella Na'geli
Navicula obsidialis Hustedt
Navicula obsoleta Hustedt
Navicula occulta Krasske
Navicula oligotraphenta Lange-Bertalot et Hofmann
Navicula oppugnata Hustedt
Navicula ordinaria Hustedt
Navicula parablis Hohn et Hellerman
Navicula paramutica Bock
Navicula pelliculosa (Brebisson) Hilse
Navicula peregrina (Ehrenberg) Kutzing
Navicula peregrinopsis Lange-Bertalot et Witkowski
Navicula perminuta Grunow
Navicula phyllepta Kutzing
Navicula phylleptosoma Lange-Bertalot
Navicula porifera (Hustedt) Lange-Bertalot
Navicula porifera var. opportuna Lange-Bertalot
Navicula praeterita Hustedt
Navicula protracta var. elliptica Gallik
Navicula pseudoarvensis Hustedt
Navicula pseudobryophila Hustedt
Navicula pseudolanceolata Lange-Bertalot
Navicula pseudosilicula Hustedt
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NLA Diatom and Non-Diatom Taxa
Navicula pseudosubtilissima Manguin
Navicula pseudoventralis Hustedt
Navicula pupula van. aquaductae (Krasske) Hustedt
Navicula pupula van. nyassensis (Mullen) Lange-Bertalot
Navicula radiosa Kutzing
Navicula radiosafallax Lange-Bertalot
Navicula radiosiola Lange-Bertalot
Navicula recens Lange-Bertalot
Navicula regularis Hustedt
Navicula reichardtiana Lange-Bertalot
Navicula reichardtiana var. crassa Lange-Bertalot et Hofmann
Navicula reinhardtii (Grunow) Grunow
Navicula rhynchocephala Kutzing
Navicula rhynchocephala var. amphiceros (Kutzing) Grunow
Navicula riediana Lange-Bertalot et Rumrich
Navicula rostellata Kutzing
Navicula rotunda Hustedt
Navicula rotundata Hantzsch
Navicula ruttneri var. rostrata Hustedt
Navicula salinarum Grunow
Navicula salinicola Hustedt
Navicula schadei Krasske
Navicula schmassmannii Hustedt
Navicula schroeterii Meister
Navicula schroeterii Meister
Navicula secreta var. apiculata Patrick
Navicula semen Ehrenberg
Navicula seminuloides Hustedt
Navicula splendicula VanLandingham
Navicula spp.
Navicula stankovicii Hustedt
Navicula striolata (Grunow) Lange-Bertalot in Krammer et Lange-Bertalot
Navicula stroemii Hustedt
Navicula subadnata Hustedt
Navicula subinflata Grunow
Navicula subinflatoides Hustedt
Navicula sublucidula Hustedt
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NLA Diatom and Non-Diatom Taxa
Navicula subminuscula Manguin
Navicula submuralis Hustedt
Navicula subplacentula Hustedt
Navicula subrhynchocephala Hustedt
Navicula subrhyncocephala Hustedt
Navicula subrotundata Hustedt
Navicula suecorum van. dismutica (Hustedt) Lange-Bertalot
Navicula symmetrica Patrick
Navicula tenelloides Hustedt
Navicula tridentula Krasske
Navicula tripunctata (Mullen) Bory
Navicula trivialis Lange-Bertalot
Navicula trivialis var. oligotraphenta Lange-Bertalot et Hofmann
Navicula trophicatrix Lange-Bertalot in Lange-Bertalot et Metzeltin
Navicula upsaliensis (Grunow) Peragallo
Navicula utermoehlii Hustedt
Navicula vaucheriae Petersen
Navicula veneta Kutzing
Navicula ventralis Krasske
Navicula viridula (Kutzing) Kutzing emend. Van Heurck
Navicula viridulacalcis (Hustedt) Lange-Bertalot
Navicula vitabunda Hustedt
Navicula vulpina Kutzing
Navicula wiesneri Lange-Bertalot
Navicula wildii Lange-Bertalot
Navicula witkowskii Lange-Bertalot, Iserentant et Metzeltin
Naviculadicta elorantana Lange-Bertalot in Lange-Bertalot et Metzeltin
Naviculadicta raederiae Lange-Bertalot in Lange-Bertalot et Metzeltin
Neidium affine (Ehrenberg) Pfitzer
Neidium alpinum Hustedt
Neidium amphigomphus (Ehrenberg) Pfitzer
Neidium ampliatum (Ehrenberg) Krammer
Neidium apiculatum Reimer
Neidium binodeformis Krammer
Neidium binodis (Ehrenberg) Hustedt
Neidium bisulcatum (Lagerstedt) Cleve
Neidium calvum 0strup
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NLA Diatom and Non-Diatom Taxa
Neidium capsulare Thomas et Kociolek
Neidium carterii Krammer
Neidium densestriatum (0strup) Krammer
Neidium dilatatum (Ehrenberg) Cleve
Neidium dubium (Ehrenberg) Cleve
Neidium excisum Krammer et Metzeltin
Neidium hercynicum Mayer
Neidium hitchcockii (Ehrenberg) Cleve
Neidium iridis (Ehrenberg) Cleve
Neidium iridis var. subundulatum (Cleve-Euler) Reimer
Neidium ladogensis (Cleve) Foged
Neidium productum (Smith) Cleve
Neidium septentrionale Cleve-Euler
Neidium spp.
Neidium stoermerii Thomas et Kociolek
Neidium temperei Reimer
Nitzschia acicularioides Hustedt
Nitzschia acicularis (Kutzing) Smith
Nitzschia acidoclinata Lange-Bertalot
Nitzschia acula Hantzsch
Nitzschia aequorea Hustedt
Nitzschia agnita Hustedt
Nitzschia alpina Hustedt
Nitzschia amphibia Grunow
Nitzschia angusta (Gregory)
Nitzschia angustata (Smith) Grunow
Nitzschia angustatula Lange-Bertalot
Nitzschia angustiforaminata Lange-Bertalot
Nitzschia archibaldii Lange-Bertalot
Nitzschia aurariae Cholnoky
Nitzschia bacilliformis Hustedt
Nitzschia bacillum Hustedt
Nitzschia behrei Hustedt
Nitzschia biacrula Hohn et Hellerman
Nitzschia bita Hohn et Hellerman
Nitzschia bremensis Hustedt
Nitzschia brevissima Grunow ex Van Heurck
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NLA Diatom and Non-Diatom Taxa
Nitzschia bryophila (Hustedt) Hustedt
Nitzschia bulnheimiana Rabenhorst
Nitzschia calcicola Aleem et Hustedt
Nitzschia capitellata Hustedt
Nitzschia circumsuta (Bailey) Grunow
Nitzschia clausii Hantzsch
Nitzschia communis Rabenhorst
Nitzschia commutata Grunow
Nitzschia commutatoides Lange-Bertalot
Nitzschia compressa var. balatonis Lange-Bertalot
Nitzschia compressa var. minor H.L. Smith ex Boyer
Nitzschia desertorum Hustedt
Nitzschia dissipata (Kutzing) Grunow
Nitzschia dissipata var. media (Hantzsch) Grunow
Nitzschia dissipata var. undulata Sovereign
Nitzschia distans Gregory
Nitzschia diversa Hustedt
Nitzschia draveillensis Coste et Ricard
Nitzschia dubia Smith
Nitzschia elegantula Grunow
Nitzschia filiformis (Smith) Van Heurck
Nitzschia flexa Schumann
Nitzschia flexoides Geitler
Nitzschia fonticola Grunow
Nitzschia fonticola var. pelagica Hustedt
Nitzschia fossilis Grunow
Nitzschia frustulum (Kutzing) Grunow
Nitzschia gessneri Hustedt
Nitzschia gisela Lange-Bertalot
Nitzschia gracilis Hantzsch ex Rabenhorst
Nitzschia gracilliformis Lange-Bertalot et Simonsen
Nitzschia graciloides Hustedt
Nitzschia hantzschiana Rabenhorst
Nitzschia harder! Hustedt
Nitzschia heufleriana Grunow
Nitzschia homburgiensis Lange-Bertalot
Nitzschia hybrida Grunow
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Nitzschia hybridaeformis Hustedt
Nitzschia incognita Legler et Krasske
Nitzschia inconspicua Grunow
Nitzschia intermedia Hantzsch ex Cleve et Grunow
Nitzschia lacunarum Hustedt
Nitzschia lacuum Lange-Bertalot
Nitzschia lanceolata fo. minima Grunow
Nitzschia lanceolata Smith
Nitzschia lanceolata var. minutula Grunow
Nitzschia leistikowii Lange-Bertalot
Nitzschia levidensis var. salinarum Grunow
Nitzschia liebethruthii Rabenhorst
Nitzschia linearis (Agardh ex Smith) Smith
Nitzschia linearis var. tenuis (Smith) Grunow ex Cleve et Grunow
Nitzschia lorenziana Grunow
Nitzschia macilenta Gregory
Nitzschia microcephala Grunow
Nitzschia modesta Hustedt
Nitzschia nana Grunow ex Van Heurck
Nitzschia obtusa Smith
Nitzschia palea (Kutzing) Smith
Nitzschia paleacea Grunow ex Van Heurck
Nitzschia paleaeformis Hustedt
Nitzschia parvula Smith
Nitzschia pellucida Grunow
Nitzschia perminuta (Grunow) Peragallo
Nitzschia perspicua Cholnoky
Nitzschia pseudofonticola Hustedt
Nitzschia pumila Hustedt
Nitzschia pura Hustedt
Nitzschia pusilla Grunow
Nitzschia radicula Hustedt
Nitzschia recta Hantzsch ex Rabenhorst
Nitzschia reversa Smith
Nitzschia romana Grunow
Nitzschia rosenstockii Lange-Bertalot
Nitzschia rostellata Hustedt
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Nitzschia scalpelliformis Grunow
Nitzschia sigma (Kutzing) Smith
Nitzschia sigmoidea (Nitzsch) Ehrenberg
Nitzschia silicula Hustedt
Nitzschia siliqua Archibald
Nitzschia sinuata (Thwaites) Grunow
Nitzschia sinuata var. delognei (Grunow) Lange-Bertalot
Nitzschia sinuata var. tabellaria (Grunow) Grunow
Nitzschia sociabilis Hustedt
Nitzschia solita Hustedt
Nitzschia spp.
Nitzschia subacicularis Hustedt
Nitzschia subconstricta Grunow
Nitzschia sublinearis Hustedt
Nitzschia subrostrata Hustedt
Nitzschia subtilioides Hustedt
Nitzschia subtilis Grunow
Nitzschia suchlandtii Hustedt
Nitzschia supralitorea Lange-Bertalot
Nitzschia terrestris (Petersen) Hustedt
Nitzschia thermaloides Hustedt
Nitzschia tropica Hustedt
Nitzschia tubicola Grunow in Cleve et Grunow
Nitzschia umbonata Lange-Bertalot
Nitzschia valdecostata Lange-Bertalot et Simonsen
Nitzschia valdestriata Aleem et Hustedt
Nitzschia vermicularis (Kutzing) Hantzsch in Rabenhorst
Nitzschia vitrea Norman
Nitzschia vitrea var. salinarum Grunow
Nitzschia wuellerstorffii Lange-Bertalot
Nupela Carolina Potapova et Clason
Nupela fennica (Hustedt) Lange-Bertalot
Nupela impexiformis (Lange-Bertalot) Lange-Bertalot
Nupela lapidosa (Krasske) Lange-Bertalot
Nupela neglecta Ponader, Lowe et Potapova
Nupela neotropica Lange-Bertalot
Nupela spp.
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Nupela vitiosa (Schimanski) Siver et Hamilton
Nupela wellneri (Lange-Bertalot) Lange-Bertalot
Opephora martyi Heribaud
Opephora olsenii M0ller
Opephora schwartzii (Grunow) Petit
Orthoseira dentroteres (Ehrenberg) Crawford
Orthoseira roeseana (Rabenhorst) O'Meara
Parlibellus crucicula (Smith) Witkowski, Lange-Bertalot et Metzeltin
Parlibellus protracta (Grunow) Witkowski, Lange-Bertalot et Metzeltin
Peronia fibula (Brebisson ex Kutzing) Ross
Pinnuavis elegans (Smith) Okuno
Pinnularia abaujensis (Pantocsek) Ross
Pinnularia acidophila Hofmann et Krammer
Pinnularia acoricola Hustedt
Pinnularia acrosphaeria (Brebisson) Smith
Pinnularia acrosphaeria var. turgidula Grunow ex Cleve
Pinnularia aestuarii Cleve
Pinnularia angusta (Cleve) Krammer
Pinnularia appendiculata (Agardh) Cleve
Pinnularia biceps Gregory
Pinnularia biceps var. pusilla Charles et Camburn
Pinnularia bihastata (Mann) Patrick
Pinnularia borealis Ehrenberg
Pinnularia borealis var. rectangularis Carlson
Pinnularia borealis var. scalaris (Ehrenberg) Rabenhorst
Pinnularia brandelii Cleve
Pinnularia braunii (Grunow) Cleve
Pinnularia braunii var. amphicephala fo. subconica Venkataraman
Pinnularia brebissonii (Kutzing) Rabenhorst
Pinnularia brevicostata Cleve
Pinnularia cleveii Patrick
Pinnularia dactylus Ehrenberg
Pinnularia divergens Smith
Pinnularia divergentissima (Grunow) Cleve
Pinnularia divergentissima var. subrostrata Cleve-Euler
Pinnularia formica (Ehrenberg) Patrick
Pinnularia gibba Ehrenberg
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Pinnularia gigas Ehrenberg
Pinnularia globiceps Gregory
Pinnularia hemiptera (Kutzing) Rabenhorst
Pinnularia hemiptera var. bielawskii (Heribaud et M. Peragallo in Heribaud) Cleve-Euler
Pinnularia hilseana Janisch
Pinnularia ignobilis (Krasske) Cleve-Euler
Pinnularia intermedia (Lagerstedt) Cleve
Pinnularia interrupta Smith
Pinnularia krammeri Metzeltin in Lange-Bertalot et Metzeltin
Pinnularia lagerstedtii (Cleve) Cleve-Euler
Pinnularia lapponica Hustedt
Pinnularia legumen (Ehrenberg) Ehrenberg
Pinnularia lundii Hustedt
Pinnularia macilenta (Ehrenberg) Ehrenberg
Pinnularia maior (Kutzing) Rabenhorst
Pinnularia mesogongyla Ehrenberg
Pinnularia mesolepta (Ehrenberg) Smith
Pinnularia mesolepta (Ehrenberg) Smith
Pinnularia microstauron (Ehrenberg) Cleve
Pinnularia molaris (Grunow) Cleve
Pinnularia nobilis (Ehrenberg) Ehrenberg
Pinnularia nodosa (Ehrenberg) Smith
Pinnularia nodosa var. percapitata Krammer
Pinnularia obscura Krasske
Pinnularia paulensis Grunow ex Cleve
Pinnularia pogoii Scherer
Pinnularia polyonca (Brebisson) Smith
Pinnularia renata Krammer
Pinnularia rupestris Hantzsch
Pinnularia similis Hustedt
Pinnularia sinistra Krammer
Pinnularia sistassa Carter
Pinnularia spp.
Pinnularia stomatophora (Grunow) Cleve
Pinnularia streptoraphe Cleve
Pinnularia subcapitata Gregory
Pinnularia subcommutata var. nonfasciata Krammer
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Pinnularia subgibba Krammer
Pinnularia sublinearis Grunow
Pinnularia subrostrata (Cleve) Cleve-Euler
Pinnularia sudetica (Hilse) Peragallo
Pinnularia superdivergentissima Chaumont et Germain
Pinnularia umbrosa Sovereign
Pinnularia viridiformis Krammer
Pinnularia viridis (Nitzsch) Ehrenberg
Placoneis abiskoensis (Hustedt) Lange-Bertalot et Metzeltin
Placoneis anglica (Ralfs in Pritchard) R. Lowe in J.R. Johansen et al.
Placoneis dementis (Grunow) Cox
Placoneis clementoides (Hustedt) Cox
Placoneis elginensis (Gregory) Cox
Placoneis exigua (Gregory) Mereschkowsky
Placoneis explanata (Hustedt) Cox
Placoneis gastrum (Ehrenberg) Mereschkowsky
Placoneis pseudanglica (Lange-Bertalot) Cox
Placoneis spp.
Plagiotropis lepidoptera (Gregory) Kuntze
Plagiotropis lepidoptera var. proboscidea (Cleve) Reimer
Plagiotropis vitrea (Smith) Cleve et Moller
Planothidium apiculatum (Patrick) Lange-Bertalot
Planothidium biporomum (Hohn et Hellerman) Lange-Bertalot
Planothidium calcar (Cleve) Bukhtiyarova et Round
Planothidium daui (Foged) Lange-Bertalot
Planothidium delicatulum (Kutzing) Round et Bukhtiyarova
Planothidium distinctum (Messikommer) Lange-Bertalot
Planothidium fossile (Tempere et Peragallo) Lowe
Planothidium frequentissimum (Lange-Bertalot) Lange-Bertalot
Planothidium hauckianum (Grunow) Round et Bukhtiyarova
Planothidium haynaldii (Schaarschmidt) Lange-Bertalot
Planothidium joursacense (Heribaud) Lange-Bertalot
Planothidium lanceolatum (Brebisson) Lange-Bertalot
Planothidium minutissimum (Krasske) Lange-Bertalot
Planothidium oestrupii (Cleve-Euler) Round et Bukhtiyarova
Planothidium peragalli Brun et Heribaud
Planothidium peragalli var. parvulum (Patrick) Andresen, Stoermer et Kreis
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Planothidium robustius (Hustedt) Lange-Bertalot
Planothidium spp.
Planothidium stewartii (Patrick) Lange-Bertalot
Planothidium ziegleri (Lange-Bertalot) Lange-Bertalot
Platessa conspicua (Mayer) Lange-Bertalot
Platessa holsatica (Hustedt) Lange-Bertalot
Platessa hustedtii (Krasske) Lange-Bertalot
Pleurosigma angulatum (Quekett) Smith
Pleurosigma delicatulum Smith
Pleurosigma elongatum Smith
Pleurosigma salinarum Grunow
Pleurosigma spp.
Pleurosira laevis (Ehrenberg) Compere
Psammodictyon constrictum (Gregory) Mann
Psammodictyon panduriforme (Gregory) Mann
Psammothidium abundans fo. rosenstockii Lange-Bertalot ex Lange-Bertalot
Psammothidium altaicum Bukhtiyarova
Psammothidium bioretii (Germain) Bukhtiyarova et Round
Psammothidium chlidanos (Hohn et Hellerman) Lange-Bertalot
Psammothidium didymum (Hustedt) Bukhtiyarova et Round
Psammothidium grischunum (Wuthrich) Bukhtiyarova et Round
Psammothidium helveticum (Hustedt) Bukhtiyarova et Round
Psammothidium kryophilum (Petersen) Reichardt
Psammothidium kuelbsii (Lange-Bertalot et Krammer) Bukhtiyarova et Round
Psammothidium lacus-vulcani (Lange-Bertalot et Krammer) Bukhtiyarova et Round
Psammothidium lauenburgianum (Hustedt) Bukhtiyarova et Round
Psammothidium levanderi (Hustedt) Czarnecki
Psammothidium marginulatum (Grunow) Bukhtiyarova et Round
Psammothidium rossii (Hustedt) Bukhtiyarova et Round
Psammothidium sacculum (Carter) Bukhtiyarova
Psammothidium scoticum (Flower et Jones) Bukhtiyarova et Round
Psammothidium spp.
Psammothidium subatomoides (Hustedt) Bukhtiyarova et Round
Psammothidium ventralis (Krasske) Bukhtiyarova et Round
Pseudostaurosira brevistriata (Grunow) Williams et Round
Pseudostaurosira clavatum Morales
Pseudostaurosira parasitica (Smith) Morales
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Pseudostaurosira parasitica van. subconstricta (Grunow) Morales
Pseudostaurosira polonica Morales et Edlund
Pseudostaurosira pseudoconstruens (Marciniak) Williams et Round
Pseudostaurosira robusta (Fusey) Williams et Round
Pseudostaurosira spp.
Pseudostaurosira subsalina (Hustedt) Morales
Pseudostaurosira trainorii Morales
Pseudostaurosira zeilleri (Heribaud) Williams et Round
Pseudostaurosiropsis spp.
Pseudostaurosiropsis spp.
Puncticulata bodanica (Grunow in Schneider) Hakansson
Puncticulata comta (Ehrenberg) Hakansson
Puncticulata radiosa (Lemmermann) Hakansson
Puncticulata spp.
Reimeria sinuata (Gregory) Kociolek et Stoermer
Reimeria uniseriata Sala, Guerrero et Ferrario
Rhizosolenia eriensis Smith
Rhoicosphenia abbreviata (Agardh) Lange-Bertalot
Rhoicosphenia spp.
Rhopalodia acuminata Krammer
Rhopalodia brebissonii Krammer
Rhopalodia gibba (Ehrenberg) Miiller
Rhopalodia gibba var. minuta Krammer
Rhopalodia gibberula (Ehrenberg) Miiller
Rhopalodia musculus (Kutzing) Miiller
Rhopalodia operculata (Agardh) Hakansson
Rhopalodia rupestris (Smith) Krammer
Rhopalodia spp.
Rossithidium duthii (Sreenivasa) Kingston
Rossithidium linearis (Smith) Round et Bukhtiyarova
Rossithidium petersennii (Hustedt) Round et Bukhtiyarova
Rossithidium pusillum (Grunow) Round et Bukhtiyarova
Scoliotropis latestriata (Brebisson in Kutzing) Cleve
Sellaphora alastos (Hohn et Hellerman) Lange-Bertalot et Metzeltin
Sellaphora americana (Ehrenberg) Mann
Sellaphora bacillum (Ehrenberg) Mann
Sellaphora disjuncta (Hustedt) Mann
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Sellaphora hustedtii (Krasske) Lange-Bertalot
Sellaphora laevissima (Kutzing) Mann
Sellaphora mutata (Krasske) Lange-Bertalot
Sellaphora pupula (Kutzing) Meresckowsky
Sellaphora seminulum (Grunow) Mann
Sellaphora spp.
Simonsenia delognei (Grunow) Lange-Bertalot
Skeletonema costatum (Greville) Cleve
Skeletonema potamos (Weber) Hasle
Stauroforma exiguiformis (Lange-Bertalot) Flower, Jones et Round
Stauroneis acuta Smith
Stauroneis agrestis Petersen
Stauroneis alpina Hustedt
Stauroneis anceps Ehrenberg
Stauroneis borrichii (Petersen) Lund
Stauroneis dubitabilis Hustedt
Stauroneis fluminea Patrick et Freese
Stauroneis gracilis Ehrenberg
Stauroneis gracillima Hustedt
Stauroneis kriegeri Patrick
Stauroneis legumen (Ehrenberg) Kutzing
Stauroneis livingstonii Reimer
Stauroneis nebulosa (Krasske) Lange-Bertalot in Lange-Bertalot, Kulbs, Lauser, Norpel-Schempp et
Willmann
Stauroneis neohyalina Lange-Bertalot et Krammer in Lange-Bertalot et Metzeltin
Stauroneis nobilis Schumann
Stauroneis obtusa Lagerstedt
Stauroneis phoenicenteron (Nitzsch) Ehrenberg
Stauroneis phoenicenteron (Nitzsch) Ehrenberg
Stauroneis producta Grunow
Stauroneis prominula (Grunow) Hustedt
Stauroneis recondita Krasske
Stauroneis schimanskii Krammer
Stauroneis siberica (Grunow in Cleve et Grunow) Lange-Bertalot et Krammer in Lange-Bertalot et
Metzeltin
Stauroneis smithii Grunow
Stauroneis spp.
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Stauroneis staurolineata Reimer
Stauroneis stodderi Lewis
Stauroneis stodderi Lewis
Stauroneis tackei Krammeret Lange-Bertalot
Stauroneis thermicola (Petersen) Lund
Staurosira bidens (Heiberg) Grunow
Staurosira construens Ehrenberg
Staurosira construens var. venter (Ehrenberg) Hamilton
Staurosira spp.
Staurosirella berolinensis (Lemmermann) Bukhtiyarova
Staurosirella lapponica (Grunow) Williams et Round
Staurosirella leptostauron (Ehrenberg) Williams et Round
Staurosirella oldenburgiana (Hustedt) Morales
Staurosirella pinnata (Ehrenberg) Williams et Round
Staurosirella spp.
Staurosirella subrobusta Morales et Manoylov
Stenopterobia anceps (Lewis) Brebisson ex Van Heurck
Stenopterobia curvula (Smith) Krammer
Stenopterobia delicatissma (Lewis) Brebisson
Stenopterobia densestriata (Hustedt) Krammer
Stenopterobia planctonica Metzeltin et Lange-Bertalot
Stenopterobia spp.
Stephanodiscus agassizensis Hakansson et Kling
Stephanodiscus agassizensis Hakansson et Kling
Stephanodiscus alpinus Hustedt
Stephanodiscus astraea (Ehrenberg) Grunow
Stephanodiscus binderanus (Kutzing) Krieger
Stephanodiscus binderanus var. oestrupii Cleve-Euler
Stephanodiscus carconensis Grunow
Stephanodiscus excentricus Hustedt emend Hakansson et Stoermer
Stephanodiscus hantzschii Grunow
Stephanodiscus medius Hakansson
Stephanodiscus minutulus (Kutzing) Cleve et Moller
Stephanodiscus neoastraea Hakansson et Hickel
Stephanodiscus niagarae Ehrenberg
Stephanodiscus rotula (Kutzing) Hendey
Stephanodiscus spp.
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Stephanodiscus subtranssylvanicus Gasse
Stephanodiscus vestibulis Hakansson, Theriot et Stoermer
Stephanodiscus yellowstonensis Theriot et Stoermer
Surirella amphioxys Smith
Surirella angusta Kutzing
Surirella bifrons Ehrenberg
Surirella birostrata Mayer
Surirella biseriata Brebisson
Surirella bohemica Maly
Surirella brebissonii Krammeret Lange-Bertalot
Surirella brebissonii var. kuetzingii Krammer et Lange-Bertalot
Surirella brebissonii var. punctata Krammer
Surirella brightwellii Smith
Surirella capronii Brebisson
Surirella constricta Smith
Surirella crumena Brebisson
Surirella didyma Kutzing
Surirella elegans Ehrenberg
Surirella gracilis (Smith) Grunow
Surirella guatimalensis Ehrenberg
Surirella helvetica Brun
Surirella helvetica Brun
Surirella hoefleri Hustedt
Surirella linearis Smith
Surirella linearis var. constricta Grunow
Surirella minuta Brebisson
Surirella minuta Brebisson
Surirella ovalis Brebisson
Surirella ovalis Brebisson
Surirella patella Kutzing
Surirella peisonis Pantocsek
Surirella roba Leclercq
Surirella robusta Ehrenberg
Surirella splendida (Ehrenberg) Kutzing
Surirella spp.
Surirella stalagma Hohn et Hellerman
Surirella striatula Turpin
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Surirella subsalsa van. minuta Smith
Surirella tenera Gregory
Surirella tenuis Mayer
Surirella utahensis (Grunow in Schmidt et al.) Hanna et Grant
Surirella venusta 0strup
Surirella visurgis Hustedt
Synedra acus Kutzing
Synedra acus Kutzing
Synedra biceps Kutzing
Synedra capitata Ehrenberg
Synedra cyclopum Brutschy
Synedra delicatissima Smith
Synedra delicatissima Smith
Synedra demerarae Grunow
Synedra dilatata Brebisson
Synedra filiformis Grunow
Synedra goulardi Brebisson
Synedra incisa Boyer
Synedra mazamaensis Sovereign
Synedra rumpens Kutzing
Synedra rumpens Kutzing
Synedra socia Wallace
Synedra spp.
Synedra ulna (Nitzsch) Ehrenberg
[Tabellaria binalis (Ehrenberg) Grunow
JTabellaria binalis var. elliptica Flower
JTabellaria fenestrata (Lyngbye) Kutzing
Tabellaria flocculosa (Roth) Kutzing
[Tabellaria flocculosa var. linearis Koppen
jTabellaria quadriseptata Knudsen
JTabellaria spp.
JTabellaria ventricosa Kutzing
Tabularia fasciculata (Agardh) Williams et Round
I
[Tabularia tabulata (Agardh) Snoeijs
JTetracyclus glans (Ehrenberg) Mills
JTetracyclus spp.
JThalassionema nitzschioides (Grunow) Van Heurck
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NLATaxa
NLA Diatom and Non-Diatom Taxa
Thalassiosira baltica (Grunow) Ostenfeld
I
[Thalassiosira bramaputrae (Ehrenberg) Hakansson et Locker
I
[Thalassiosira oestrupii (Ostenfeld) Hasle
JThalassiosira proschkinae Makarova
JThalassiosira pseudonana Hasle et Heimdal
Thalassiosira spp.
[Thalassiosira tenera Proshkina-Lavrenko
JThalassiosira visurgis Hustedt
JThalassiosira visurgis Hustedt
Thalassiosira weissflogii (Grunow) Fryxell et Hasle
.
[Tryblionella acuminata Smith
I
[Tryblionella aerophila (Hustedt) Mann
JTryblionella calida (Grunow in Cleve et Grunow) Mann
JTryblionella coarctata (Grunow) Mann
Tryblionella compressa (Bailey) Poulin
I ~
[Tryblionella constricta (Kutzing) Ralfs
JTryblionella debilis Arnott
JTryblionella gracilis Smith
Tryblionella hungarica (Grunow) Mann
[Tryblionella levidensis Smith
JTryblionella levidensis Smith
JTryblionella littoralis (Grunow) Mann
JTryblionella peisonis (Pantocsek) F.W. Mills
Tryblionella plana (Smith) Pelletan
I
[Tryblionella scalaris (Ehrenberg) Siver et Hamilton
JTryblionella spp.
JTryblionella umbilicata (Hustedt) Mann
Tryblionella victoriae Grunow
Urosolenia longiseta (Zacharias) Bukhtiyarova
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Appendix C
Supplementary Material for Vegetation—Lists of Floristic Resources
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Region I
Ballard, B.D, H.L. Whittier, C.A. Newark. 2004. Northeastern Shrub and Short Tree
Identification. SUNY. 120 pp.
Brown, L. 1992.Grasses: An Identification Guide. Houghton Mifflin Harcourt. 256 pp.
Campbell, C.S. 1978. Winter Keys to Woody Plants of Maine. University of Maine Press. 52 pp.
Common Wetland Delineation Sedges of the Northeast - Robert Lichvar
Cope. 2001. Muenscher's Keys to Woody Plants. An Expanded Guide to Native and Cultivated
Species. Comstock Publishing Associates. Cornell University Press. Ithaca. 368 pp.
Crow and Hellquist. 2005. Aquatic and Wetland plants of Northeastern North America. The
University of Wisconsin Press. Vol. 1. Pteridophytes, Gymnosperms, and Angiosperms:
Dicotyledons.; Vol. 2. Angiosperms: Monocotyledons. Madison. 400 pp.
Farnsworth, A., B. Cobb, and C. Lowe. 2005. Peterson Field Guide to Ferns, Second Edition:
Northeastern and Central North America. Houghton Mifflin Harcourt. 440 pp.
Fernald. 1950. Gray's Manual of Botany, 8th Ed. D. Van Nostrand Company, New York. 1632
pp.
Flora of North America (FNA) Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+vols. New York and Oxford. Available online:
http://hua.huh.harvard.edu/FNA/volumes.shtml and
http://www.efloras.org/flora page.aspx?flora id=1
Gleason, H.A. 1968. The new Britton and Brown illustrated flora of the Northeastern United
States and adjacent Canada. Macmillan Pub Co. New York Botanical Garden. 1098 pp.
Gleason and Cronquist. 1963. Manual of the Vascular Plants of Northeastern United States and
Adjacent Canada. Van Nostrand, Princeton NJ. 910 pp.
Gleason and Cronquist. 1991. Manual of Vascular Plants of the Northeastern United States and
Adjacent Canada. Second Edition. The New York Botanical Garden. New York. 993 pp.
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Gleason, H.A., N.H. and P.K. Holmgren. 1998. Illustrated Companion to Gleason and Cronquit's
Manual: Illustrations of the Vascular Plants of Northeastern United States and Adjacent
Canada. New York Botanical Garden, New York. 937 pp.
Haines and Vining. 1998 Flora of Maine: A Manual for Identification of Native and Naturalized
Plants of Maine. V. F. Thomas Company. Bar Harbor. 847 pp.
Haines. 2011. Flora Novae Angliae: A Manual for the Identification of Native and Naturalized
Higher Vascular Plants of New England. New England Wildflower Society. Yale
University Press. New Haven. 1008 pp.
Hallowell, A.C, and E.G. Hallowell. 2001. Fern Finder: A Guide to Native Ferns of Central and
Northeastern United States and Eastern Canada. Wilderness Press. 64 pp.
Holmgren. 1998. Illustrated Companion to Gleason and Cronquist's Manual of Vascular Plants.
The New York Botanical Garden. New York. 919 pp.
Knobel, E. 1977. Field Guide to the Grasses, Sedges, and Rushes of the United States. Dover
Publications. 96 pp.
Magee and Ahles. 2007. Flora of the Northeast. A Manual of the Vascular Flora of New
England and Adjacent New York. Second edition with companion CD. University of
Massachusetts Press. Amherst. 1214 pp.
McQueen, C.B. 1990. Field Guide to the Peat Mosses of Boreal North America. University
Press of New England. 163 pp.
Michigan Flora Online. A. A. Reznicek, E. G. Voss, & B. S. Walters. February 2011. University
of Michigan. Web. 6-17-2011. http://michiganflora.net/home.aspx.
Newcomb. 1989. Newcomb's Wildflower Guide. Little, Brown and Company. New York. 490 pp.
Peterson, R.T. 1968. A Field Guide to Wildflowers of Northeastern and North-Central North
America. Houghton Mifflin, Boston. 420 pp.
Petrides, G.A and R.T. Peterson. 1973. A Field Guide to Trees and Shrubs: Northeastern and
North-Central United States and Southeastern and South-Central Canada. 464 pp.
Rhoads and Block. 2007. The Plants of Pennsylvania. An Illustrated Manual. Second Edition.
Philadelphia. 1056 pp.
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Sample, J. 1999. The Goldenrods of Ontario: Solidago L. and Euthamia Nutt. 3rd Edition.
University of Waterloo Biology Department. 90 pp.
Standley. 2011. Field Guide to Carex of New England. Special Publication of the New England
Botanical Club. Cambridge. 182.pp.
Voss. Michigan Flora. A Guide to the Identification and Occurrence of the Native and
Naturalized Seed-plants of the State; Part I: Gymnosperms and Monocots. Cranbrook
Institute of Science Bulletin 55 & University of Michigan Herbarium, xv + 488 pp. 1972.;
Part II: Dicots (Saururaceae-Cornaceae). Cranbrook Institute of Science Bulletin 59 &
University of Michigan Herbarium, xix + 724 pp. 1985; Part III: Dicots (Pyrolaceae-
Compositae). Cranbrook Institute of Science Bulletin 61 & University of Michigan
Herbarium, xii + 622 pp. 1996. Bloomfield Hills.
Weldy, Troy and David Werier. 2011 New York Flora Atlas. [S. M. Landry and K. N. Campbell
(original application development), Florida Center for Community Design and Research.
University of South Florida]. New York Flora Association, Albany, New York.
http://newyork.plantatlas.usf.edu/
Region
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
Gleason and Cronquist. 1963. Manual of the Vascular Plants of Northeastern United States and
Adjacent Canada. Van Nostrand, Princeton NJ. 910 pp.
Gleason, H.A., N.H. and P.K. Holmgren. 1998. Illustrated Companion to Gleason and Cronquit's
Manual: Illustrations of the Vascular Plants of Northeastern United States and Adjacent
Canada. New York Botanical Garden, New York. 937pp.
Peterson, R.T. 1968. A Field Guide to Wildflowers of Northeastern and North-Central North
America. Houghton Mifflin, Boston. 420 pp.
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Region III
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
Gleason and Cronquist. 1963. Manual of the Vascular Plants of Northeastern United States and
Adjacent Canada. Van Nostrand, Princeton NJ. 910 pp.
Gleason, H.A., N.H. and P.K. Holmgren. 1998. Illustrated Companion to Gleason and Cronquit's
Manual: Illustrations of the Vascular Plants of Northeastern United States and Adjacent
Canada. New York Botanical Garden, New York. 937pp.
Peterson, R.T. 1968. A Field Guide to Wildflowers of Northeaster and North-Central North
America. Houghton Mifflin, Boston. 420 pp.
Strausbaugh, P. D. and E. L Core. 1978. Flora of West Virginia, Second Edition. Seneca
Books, Morgantown. 1079 pp.
Weakley, A. S. 2011. Flora of the Southern and Mid-Atlantic States. Working Draft of 15 May
2011. University of North Carolina Herbarium, North Carolina Botanical Garden. Chapel
Hill, NC. 1072 pp.
Region IV
Eastman, J. 1995. The book of swamp and bog. Stackpole Books, Mechanicsburg, PA.
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
Foote, L.E., S. B. Jones, Jr. 2005. Native shrubs and woody vines of the Southeast. Timber
Press, OR.
Godfrey, R.K. 1989. Trees, Shrubs, and Woody Vines of Northern Florida and Adjacent Georgia
and Alabama. The University of Georgia Press
(http://www.ugapress.uga.edu/index.php/ugapressbook/trees shrubs/)
Godfrey, R.K., J.W. Wooten. 1979. Aquatic and Wetland Plants of Southeastern United States,
Monocotyledons. The University of Georgia Press
(http://www.ugapress.uga.edu/index.php/ugapressbook/aquatic_and_wetland_plants/)
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Godfrey, R.K., J.W. Wooten. 1981. Aquatic and Wetland Plants of Southeastern United States,
Dicotyledons. The University of Georgia Press
(http://www.ugapress.uga.edu/index.php/ugapressbook/aquatic and wetland plantsi/)
Harrar, E.S., J.G. Harrar. 1962. Guide to southern trees. Dover Publications, Inc., NY.\
Weakley, A. S. 2011. Flora of the Southern and Mid-Atlantic States. Working Draft of 15 May
2011. University of North Carolina Herbarium, North Carolina Botanical Garden. Chapel
Hill, NC. 1072 pp.
Florida
Clewell, A.F. 1988. Guide to the vascular plants of the Florida panhandle. University Presses of
Florida.
Langeland, K.A., K. Craddock Burks, eds. 1998. Identification and biology of non-native plants
in Florida's natural areas. University of Florida (this may all be available on-line now at
http://aquatl.ifas.ufl.edu/)
Osorio, R. 2001. A gardeners guide to Florida's native plants. University Presses of Florida.
Tobe, J.D., K. Craddock Burks, R.W. Cantrell, M.A. Garland, M.E. Sweeley, D.W. Hall, P.
Wallace, G. Anglin, G. Nelson, J.R. Cooper, D. Bickner, K. Gilbert, N. Aymond, K.
Greenwood, and N. Raymond. 1998. Florida wetland plants: an identification manual.
Florida Department of Environmental Protection, Tallahassee, Florida, USA.
Region V
Chadde, S.W. 2002. A Great Lakes Wetland Flora: Second Edition. Pocketflora Press. 648pp.
Crow, G.E., et al. 2006. Aquatic Wetland Plants of Northeastern North America. 536pp.
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
Gleason, Henry A. and Arthur Cronquist. 1991. Manual of Vascular Plants of the NE U.S. and
Adjacent Canada. NY Botanical Garden. Bronx, NY.
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Holmgren, Noel H. et al. 1998. Illustrated Companion to Gleason and Cronquist's Manual. NY
Botanical Garden. Bronx, NY.
Newcomb, Lawrence. 1977. Newcomb's Wildflower Guide. Little, Brown, and Co. New York.
Peterson, R.T. 1968. A Field Guide to Wildflowers of Northeaster and North-Central North
America. Houghton Mifflin, Boston. 420 pp.
Swink, Floyd and Gerould Wilhelm. 1994. Plants of the Chicago Region. 4th Ed. Indiana
Academy of Science. Indianapolis.
Michigan
Barnes, Burton V. and Warren H. Wagner. 1981, 2004. Michigan Trees. The University of
Michigan Press. Ann Arbor.
Core, Earl L. and Nelle P. Ammons. 1958. Woody Plants in Winter. The Boxwood Press.
Pacific Grove, California.
Harris, James G. and Melinda Woolf Harris, 2001. Plant Identification Terminology: An
Illustrated Glossary, 2nd Ed. Spring Lake Publishing
Mohlenbrock, Robert H. 1999. The Illustrated Flora of Illinois - Sedges: Carex. Southern
Illinois University Press. Carbondale, Illinois.
Mohlenbrock, Robert H. 2001. The Illustrated Flora of Illinois - Sedges: Cyperus to Scleria.
2nd Ed. Southern Illinois University Press. Carbondale, Illinois.
Mohlenbrock, Robert H. 1970. The Illustrated Flora of Illinois - Flowering Rush to Rushes.
Southern Illinois University Press. Carbondale, Illinois.
Semple, John C. and Gordon S. Ringius. 1992. Goldenrods of Ontario. University of Waterloo.
Waterloo, Ontario, Canada.
Semple, John C., Stephen B. Heard, and Chun Sheng Xiang. 1996. The Asters of Ontario.
University of Waterloo. Waterloo, Ontario, Canada.
Soper, James H. and Margaret L. Heimburger. 1982. Shrubs of Ontario. The Royal Ontario
Museum. Toronto.
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Steven G. Newmaster, Allan G. Harris, and Linda J. Kershaw 1997. Wetland Plants of Ontario,
Lone Pine Publishing; Edmonton, Alberta, Canada.
Symonds, George W.D. 1963. The Shrub Identification Book. William Morrow & Co. New
York.
Voss, Edward G. 1972, 1985, 1996. Michigan Flora, Part I, II, III. Cranbrook Institute of
Science and University of Michigan Herbarium. Bulletin 55.
Region VI
Correll, D.S, and M.C Johnston. 1970. Manual of vascular plants of Texas. Texas Research
Foundation, renner. 1881 pp.
Diggs, G.M Jr., B.L. Lipscomb, and R.J. O'Kennon. 1999. Botanical Research Institute of Texas,
Fort Worth, Texas. 1626 pp.
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
Stutzenbaker, Charles D. 1999. Aquatic and Wetland Plants of the Western Gulf Coast. Texas
Parks and Wildlife Press (distributed by the University of Texas Press). 465pp.
Arkansas
Smith. E.B. 1994. Keys to the Flora of Arkansas. University of Arkansas Press. 376 pp.
New Mexico
Martin, W.C. 1981. Flora of New Mexico. Lubrecht & Cramer, Limited. 3000 pp.
Region VII
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
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Missouri
Steyemark's Flora of Missouri, Volume 2. 2006. Missouri Botanical Garden Press. 1200 pp.
Yatskeivych, George. 1999. Steyermark's Flora of Missouri, Volume 1. Missouri Botanical
Garden Press. 991 pp.
Region VIM
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+vols. New York and Oxford.
Great Plains Flora Association. 1986. Atlas of the flora of the Great Plains. Coordinator, R.L.
McGregor Editor!. M. Barkley. University Press of Kansas 1408 pp.
Hitchcock, C.L and A. Cronquist. 1973. Flora of the Pacific Northwest: An Illustrated Manual.
University of Washington Press. 730 pp.
Wilson et al. 2008. Field Guide to the Sedges of the Pacific Northwest. Oregon State University
Press.
Colorado
Beidleman, L.H., R.G. Beidleman, and B.E. Willard. 2000. Plants of Rocky Mountain National
Park. Falcon Press. 266 pp.
Carter, J. L. 2006. Trees and Shrubs of Colorado, Revised and Expanded. Mimbres Publishing.
165pp.
Hurd, E.G., N.L. Shaw, J. Mastroguiseppe, L.C. Smithman, and S. Goodrich. 1998. Field Guide
to Intermountain Sedges. RMRS-GTR-10. USDA Rocky Mountain Research Station. 282
pp. (http://www.fs.fed.us/rm/pubs/rmrs gtr010.html.)
Kershaw L.J., A. MacKinnon, and J. Pojar. 1998. Plants of the Rocky Moutains. Lone Pine
Press. 384 pp.
Weber, W.A. and R.C. Wittmann. 2001. Colorado Flora Eastern Slope, Third Edition. University
Press of Colorado. 521 pp.
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Weber, W.A. and R.C. Wittmann. 2001. Colorado Flora Western Slope, Third Edition. University
Press of Colorado. 488 pp.
Montana
Dorn, R.D. and J.L Dorn. 1984. Vascular Plants of Montana. Mountain West Pub. 276 pp.
Region IX
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
California
Hickman, James C., Editor. 1993. The Jepson Manual of Higher Plants of California. University
of California Press, Berkeley, California. Third printing with corrections 1996.
Region X
Cooke, S.S. 1997. A Field Guide to the Common Wetland Plants of Western Washington &
Northwestern Oregon. Seattle Audubon Society 403 pp.
Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of
Mexico. 12+ vols. New York and Oxford.
Guard, B.J. 2010. Wetland Plants of Oregon and Washington. Lone Pine Publishing 240 pp.
Hitchcock, C.L, and A. Cronquist. 1973. Flora of the Pacific Northwest: An Illustrated Manual.
University of Washington Press 730 pp.
Jolley, R. 1988. Wildflowers of the Columbia Gorge. Oregon Historical Society Press, First
Edition 331 pp.
Kozloff, E. 2005. Plants of Western Oregon, Washington & British Columbia. Timber Press Inc.
608 pp.
Pojar, J., and A. MacKinnon. 1994. Plants of the Pacific Northwest Coast: Washington, Oregon,
British Columbia & Alaska. Lone Pine Publishing 528 pp.
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Parish, R. 1999. Plants of Southern Interior British Columbia and the Inland Northwest. Lone
Pine Publishing 464 pp.
Tuner, M. and P. Gustafson. 2006. Wildflowers of the Pacific Northwest. Timber Press, Inc512
pp.
Wilson, B., R. Brainerd, D. Lytjen, B. Newhouse, and N. Otting. 2008. Field Guide to the Sedges
of the Pacific Northwest. Oregon State University 432 pp.
http://www.okanoganl.com/botany/flora/wooten-salix-current.pdf
http://www.wsdot.wa.gov/NR/rdonlyres/5DABA8B5-10F6-4CBA-B87D-E76F814CA66
9/0/SalixKev.pdf
http://info.ag.uidaho.edu/pdf/SB/SB039.pdf
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Appendix D
Plant Pressing and Mounting
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Pressing Samples
Plants must be pressed and dried immediately upon reception at the. Immediately pressing and
drying results in the highest quality samples which facilitate identification.
Standard Procedures for Pressing Plant Samples
Samples are pressed and dried in a standard plant press (30 X 45 cm or 12 X 18 inches),
composed of:
a. A rigid, breathable wooden frame
b. Corrugated cardboard ventilators to allow air flow through the press
c. Blotter paper to absorb moisture
d. Folded newsprint to contain the plant material
e. Straps with buckles to tighten the press
Assembling the press
Each newspaper sample folder with plant material is sandwiched between 2 moisture-absorbing
blotters. The "blotter-newsprint sandwiches" are sandwiched between corrugated cardboard,
ensuring that the corrugations of the cardboard run parallel to the shorter dimension (30 cm) for
best air circulation. Bulky samples may require extra blotters and cardboard. Specifics on how
to display plant material within the newsprint folders and assemble the press are provided
below:
7. On the bottom wooden frame of the press, place cardboard and on top the cardboard add a
blotter. Lay a newsprint folder on top of the blotter.
8. Plant material will be placed inside this sheet of folded newsprint (folded size 30 X 45 cm).
Ideally, samples should not exceed this size so that they can be entirely enclosed in the
plant press and because the heavy herbarium paper on which samples may later be
mounted has dimensions of 29 X 42 cm (11.5 x 16.5 inches).
9. For every plant sample, clearly write on the outside of its newsprint folder, using water-proof
ink, the sample number (see Vegetation Chapter in FOM) and other pertinent tracking
information (plant name or pseudonym, X-Point number, collection date, and collector(s)
name(s)).
10. Clean as much dirt as possible off the plant material before placing it in the newsprint folder.
11. Open the newsprint folder and place a filled out Plant Sample Label Form (see Fig. 4-3) on
the bottom fold of the paper.
12. Carefully arrange the plant material on the bottom fold of the newsprint to display diagnostic
features.
13. Lay the plants flat and avoid overlapping plant parts.
14. Spread leaves, flowers, and fruits so they can be easily observed from different
perspectives.
15. Show upper and lower surfaces of leaves and flowers.
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16. If possible, arrange pressed material to show some flowers with the blossom open, and
some flowers and fruits are pressed in longitudinal and transverse views.
17. Multiples of smaller plants should be pressed together in order to provide ample material for
study.
18. For large samples, bend stems sharply into a V, N or W shapes so they fit within the press
frame. Do not curve or twist the stems.
19. In order to insure rapid and thorough drying, thick stems, large fruits, or bulbs may be cut in
half lengthwise. Areas around thick structure may need additional absorptive layers, e.g.,
extra newspaper or blotters.
20. Examples of small, loose plant parts, such as seeds or Carex perigynia, should be placed in
a small paper packet or envelope inside of the newspaper.
21. Once the plant material is satisfactorily arranged, fold the newsprint closed.
22. Add another blotter, then a cardboard on top of the newsprint folder. To begin pressing the
next sample, place a blotter over the top cardboard in the stack. Add a newsprint folder and
repeat steps 3-13 until the press is full or all samples are included.
23. Use two adjustable straps to firmly hold the plant press and its contents. The plant press
must be kept tight to prevent shrinkage and wrinkling of the plant material. It should not be
possible to move the blotters or cardboards from the side in a properly tightened press.
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Mounting Plant Samples
Vascular plants should:
• Be mounted on archival-quality paper measuring 11.5 X 16.5 inches
• Be mounted using commercially available acid free adhesive, such as polyvinyl acetate
(PVA)
• Allow for placement of a properly filled out label
• Have an acid free fragment envelop (where necessary for seeds and flowers)
Labeling Herbarium Sheets
Herbarium labels are considered field data, and an herbarium sheet is incomplete in the
absence of accurate label data. A completed Plant Sample Label Form (Figure 4-5) should be
affixed to the herbarium sheet in preparation for storage. Much of the information required to
complete the label Data Form is available on the Plant Sample Label Form (Vegetation
Chapters in FOM) EPA will provide Label Data Forms.
Information to be included on voucher sample labels
• Sample Number: NWCA Site Number-Plant sample number-Plot module number (if
applicable). Plant sample numbers for Samples are ordered chronologically for each site
beginning with one. For example, the Sample number for the 5th species collected at
NWCA X-Point 0762 in Plot module 3 would be NWCA-0762-005-3. For Samples not
collected in a vegetation plot, the plot module number is 0.
• Scientific name: The Genus, species, and authority are recorded.
• Collector(s) name: List the first name, middle initial, and surname of the person or
persons who collected the Sample.
• Date of collection: List the day, month (spelled out or abbreviated), and 4 digit year, e.g.,
3 May 2003, for when the collection was made.
• Identified By: This item is left blank until the Sample is identified. Upon identification, the
first name, middle initial, and surname of the person who identified the plant.
• Verified By: This item is filled in only for cases where a second person verifies the first
identification, or corrects the original identification. The first, middle initial, and surname of
the person who did the verification or made the correction. If a name correction is made, it
is noted in the Scientific Name field and initialed and dated by the verifying botanist.
• Detailed location: (note: much of this information should be available in site packets or
on other data forms - so perhaps need not be repeated here as long as it is available to
whoever is doing plant identification or to herbaria where vouchers might be deposited):
o County and state
o Public or private land and land manager or landowner information
o Description of the location in reference to roads, road junctions, mile markers and
distances from cities and/or towns, or nearby natural features.
o Latitude and longitude, section, township and range
o Global Positioning System (GPS) coordinates
o Elevation
• Habitat: The type of plant community or setting where the plant is growing, (e.g., such as
wetland type (Cowardin, NVC, HGM), wetland community type (forested wetland,
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emergent marsh, wet prairie, mountain bog, etc.), anthropogenic disturbances (urban
setting type), and, if known, other plants growing in association (associated species
information would be available from the plot.
• Plant habit: Describe key features of the plant such as growth form (tree, shrub, vine,
herb), approximate height, longevity (annual, biennial, perennial), clonal, rhizomatous,
tussock-forming, etc.
• Frequency: Indicate whether the species is rare, occasional, frequent or common at the
site, and whether based on canopy cover it is a dominant, moderate cover, or low cover
species.
• Plant description: List any characteristics of the plant which may be lost upon drying,
such as flower/fruit color and fragrance, leaf orientation and aroma.
Plants of the National Wetlands Condition Assessment
Species Name Fami|y Name
Detailed Location:
Habitat:
Plant Habit:
Plant Description:
Collector's Name: Specimen Number Date (Month, Day, Year)
Identifier's name
Figure 4-5: Format for Herbaria Labels
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