oEPA
United States
Environmental Protection
Agency
Analytic Methods for the Oil and Gas
Extraction Point Source Category
U.S. Environmental Protection Agency
Engineering and Analysis Division
Office of Water
1200 Pennsylvania Avenue, NW
Washington, D.C. 20460
December 2011
EPA-821-R-11-004
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CONTENTS
Page
1. SUMMARY OF METHODS 1-1
1.1 Static Sheen Test (EPA Method 1617) 1-5
1.2 Drilling Fluids Toxicity Test (EPA Method 1619) 1-5
1.3 Procedure for Mixing Base Fluids with Sediments (EPA Method 1646) 1-5
1.4 Protocol for the Determination of Degradation of Non-Aqueous Base
Fluids in a Marine Closed Bottle Biodegradation Test System: Modified
ISO 11734:1995 (EPA Method 1647) 1-6
1.5 Determination of Crude Oil Contamination in Non-Aqueous Drilling
Fluids by Gas Chromatography/Mass Spectrometry (GC/MS) (EPA
Method 1655) 1-7
1.6 Reverse Phase Extraction (RPE) Method for Detection of Oil
Contamination in Non-Aqueous Drilling Fluids (NAF) (EPA Method
1670) 1-7
1.7 Determination of the Amount Of Non-Aqueous Drilling Fluid (NAF) Base
Fluid from Drill Cuttings by a Retort Chamber (Derived From API
Recommended Practice 13B-2) (EPA Method 1674) 1-8
1.8 PAH Content of Oil by HPLC/UV (EPA Method 1654, Revision A) 1-8
1.9 Sediment Toxicity Test for NAF and SBM (EPA Method 1644) 1-8
1.10 Previous Publication of Oil and Gas Extraction Point Source Category
Analytic Methods 1-9
2. STATIC SHEEN TEST (EPA METHOD 1617) 2-11
2.1 Scope and Application 2-11
2.2 Summary of Method 2-11
2.3 Interferences 2-11
2.4 Apparatus, Materials, and Reagents 2-11
2.5 Calibration 2-12
2.6 Quality Control Procedures 2-12
2.7 Sample Collection and Handling 2-12
2.8 Procedure 2-13
3. DRILLING FLUIDS TOXICITY TEST (EPA METHOD 1619) 3-1
3.1 Scope and Application 3-1
3.2 Summary of Method 3-1
3.3 Sample Collection 3-1
3.4 Suspended Particulate Phase Sample Preparation 3-2
3.5 Guidance for Performing Suspended Particulate Phase Toxicity Tests
Using Mysidopsis bahia 3-4
3.6 Methods for Positive Control Tests (Reference Toxicant) 3-7
3.7 Randomization Procedure 3-7
3.8 References 3-12
4. PROCEDURE FOR MIXING BASE FLUIDS WITH SEDIMENTS (EPA METHOD 1646) 4-1
4.1 Determining the Wet to Dry Ratio for the Control Sediment 4-1
4.2 Determining the Density of the Wet Control or Dilution Sediment 4-1
4.3 Determining the Amount of Base Fluid Needed 4-1
4.4 Primary Mixing 4-2
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4.5 Testing for Homogeneity of Base Fluid 4-2
4.6 Commencing the Sediment Toxicity Test 4-2
4.7 References 4-3
5. PROTOCOL FOR THE DETERMINATION OF DEGRADATION OF NON-AQUEOUS BASE
FLUIDS IN A MARINE CLOSED BOTTLE BIODEGRADATION TEST SYSTEM: MODIFIED
ISO 11734:1995 (EPAMETHOD 1647) 5-1
5.1 Summary of Method 5-1
5.2 System Requirements 5-1
5.3 Test Set Up 5-2
5.4 Concentration Verification Chemical Analyses 5-6
5.5 Gas Monitoring Procedures 5-7
5.6 Test Acceptability and Interpretation 5-8
5.7 Methane Measurement 5-9
5.8 Concentration Verification Analysis 5-11
5.9 Program Quality Assurance and Quality Control 5-12
6. DETERMINATION OF CRUDE OIL CONTAMINATION IN NON-AQUEOUS DRILLING
FLUIDS BY GAS CHROMATOGRAPHY/MASS SPECTROMETRY (GC/MS) (EPA
METHOD 1655) 6-1
6.1 Scope and Application 6-1
6.2 Summary of Method 6-1
6.3 Definitions 6-2
6.4 Interferences and Limitations 6-2
6.5 Safety 6-2
6.6 Apparatus and Materials 6-3
6.7 Reagents and Standards 6-4
6.8 Sample Collection Preservation and Storage 6-6
6.9 Quality Control 6-6
6.10 Calibration 6-9
6.11 Procedure 6-11
6.12 Calculations 6-15
6.13 Method Performance 6-16
6.14 Pollution Prevention 6-16
6.15 Waste Management 6-16
6.16 References 6-17
6.17 Schematic Flowchart for Qualitative Identification 6-18
7. REVERSE PHASE EXTRACTION (RPE) METHOD FOR DETECTION OF OIL
CONTAMINATION IN NON-AQUEOUS DRILLING FLUIDS (NAF) (EPA METHOD 1670) 7-1
7.1 Scope and Application 7-1
7.2 Summary of Method 7-1
7.3 Definitions 7-1
7.4 Interferences 7-2
7.5 Safety 7-2
7.6 Equipment and Supplies 7-2
7.7 Reagents and Standards 7-4
7.8 Sample Collection, Preservation, and Storage 7-4
7.9 Quality Control 7-4
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7.10 Calibration and Standardization 7-7
7.11 Procedure 7-7
7.12 Data Analysis and Calculations 7-9
7.13 Method Performance 7-9
7.14 Pollution Prevention 7-9
7.15 Waste Management 7-10
7.16 References 7-10
8. DETERMINATION OF THE AMOUNT OF NON-AQUEOUS DRILLING FLUID (NAF) BASE
FLUID FROM DRILL CUTTINGS BY A RETORT CHAMBER (DERIVED FROM API
RECOMMENDED PRACTICE 13B-2) (EPA METHOD 1674) 8-1
8.1 Description 8-1
8.2 Equipment 8-1
8.3 Procedure 8-2
8.4 Calculations 8-3
8.5 Requirements for Sampling Cuttings Discharge Streams for use with this
Method 8-9
8.6 Best Management Practices (BMPs) for use with this Method 8-12
9. PAH CONTENT OF OIL BYHPLC/UV (EPA METHOD 1654, REVISION A) 9-1
9.1 Scope and Application 9-1
9.2 Summary of Method 9-1
9.3 Interferences 9-1
9.4 Safety 9-2
9.5 Apparatus and Materials 9-2
9.6 Reagents 9-4
9.7 Calibration 9-5
9.8 Quality Assurance/Quality Control 9-6
9.9 Sample Collection, Preservation, and Handling 9-9
9.10 Dilution of Oil and Extracts 9-9
9.11 High-Performance Liquid Chromatography 9-10
9.12 HPLC System and Laboratory Performance 9-11
9.13 Qualitative Identification 9-12
9.14 Quantitative Determination 9-12
9.15 Method Performance 9-13
9.16 References 9-13
10. METHOD FOR CONDUCTING A SEDIMENT TOXICIT Y TEST WITH LEPTOCHEIRUS
PLUMULOSUS AND NON-AQUEOUS DRILLING FLUIDS OR SYNTHETIC-BASED
DRILLING MUDS (EPAMETHOD 1644) 10-1
10.1 Summary of Method 10-1
10.2 Test Requirements and Materials 10-1
10.3 Procedures 10-6
10.4 Recommended Test Conditions 10-8
10.5 Biological Data 10-10
10.6 Quality Control 10-10
10.7 Reference Tests 10-11
10.8 Interpretation of Result 10-12
10.9 References 10-13
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LIST OF TABLES
Page
Table 1 -1. Methods by Waste and Pollutant, Subpart A - Offshore Subcategory 1-2
Table 1 -2. Methods by Waste and Pollutant, Subpart D - Coastal Subcategory 1-4
Table 1-3. EPA Method Numbers for Oil and Gas Extraction Point Source Category
Analytical Methods and Prior CFR References in 40 CFR 435 1-10
Table 3-1. Example of a Randomization Schedule 3-9
Table 3-2. Listing of Acute Toxicity Test Data (August 1983 to September 1983) With
Eight Generic Drilling Fluids and Mysid Shrimp 3-11
Table 3-3. Partial Toxicity Test Passing Criteria 3-12
Table 5-1. Test Acceptability Criteria 5-9
Table 6-1. Gas Chromatograph/Mass Spectrometer (GC/MS) Operation Conditions 6-9
Table 6-2. Approximate Retention Time for Compounds 6-10
Table 6-3. Recommended Ion Mass Numbers 6-13
Table 9-1. Performance Data and Method Acceptance Criteria for PAH 9-14
Table 9-2. HPLC Calibration Data 9-14
Table 10-1. Culture conditions for L. plumulosus. Conditions listed are consistent with
culture conditions specified in ASTM E 1367-92 and subsequent updates (E
1367-99) 10-3
Table 10-2. Dry food portion of the diet that has been used to successfully culture L.
plumulosus 10-4
Table 10-3. Conditions for conducting 96-hour NAF and 10-day SBM sediment toxicity
tests with L. plumulosus. Conditions listed are consistent with test conditions
specified in ASTM E 1367- 92 and subsequent updates (E 1367-99) unless
otherwise noted 10-9
Table 10-4. Test acceptability requirements for 10-day NAF and 96-hr SBM tests with L.
plumulosus. Requirements listed are consistent with those specified in ASTM
E 13 67-92 and sub sequent updates (E 1367-99) 10-11
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LIST OF FIGURES
Page
Figure 3-1. Mysid Randomization Procedure 3-8
Figure 6-1. Schematic Flowchart for Qualitative Identification 6-18
Figure 9-1. Liquid Chromatography of the Three-Component Standard and of No. 2
Diesel Oil 9-15
VI
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1. SUMMARY OF METHODS
This document is a compilation of all but one of the analytical methods necessary for
demonstrating compliance with the requirements of 40 CFR Part 435, effluent limitations
guidelines and standards for the Oil and Gas Extraction Point Source Category. These methods
are tests for regulated pollutant parameters. The waste sources and pollutant parameters
regulated in 40 CFR Part 435 are listed in the following tables along with the required analytical
or test method, and the subpart that cites the method. Table 1-1 presents the waste sources and
pollutant parameters regulated by Subpart A - Offshore Subcategory. Table 1-2 presents the
same information for Subpart D - Coastal Subcategory.
These methods were developed during two rulemakings: Offshore Subcategory (March 4, 1993;
58 FR 12454) and the Synthetic-based Drilling Fluids (SBF) Effluent Guidelines (January 22,
2001; 66 FR 6849). These analytic methods were developed in consultation and collaboration
with industry. For example, the analytic methods for the Offshore Subcategory were developed
with industry in the 1980's and included the Static Sheen Test and Drilling Fluids Toxicity Test
(see August 26, 1985; 50 FR 34596). EPA again worked very closely with industry and adopted
their recommendations in promulgating additional analytical methods for the Synthetic-based
Drilling Fluids Effluent Guidelines (see January 22, 2001; 66 FR 6893). These additional
analytical methods included tests for PAH content (as phenanthrene), sediment toxicity,
biodegradation rate, and percentage of base fluid on drill cuttings (API retort method). EPA
listed the analytic methods and in some cases re-printed the analytic methods in the Code of
Federal Regulations (CFR).
As previously noted there is one method that is not included in this document, which relates to
Cook Inlet, Alaska, operators that are regulated by the Coastal Subcategory (Subpart D). The
Coastal Subcategory prohibits the discharge non-aqueous drill cuttings unless there are technical
limitations. Appendix 1 of 40 CFR Part 435 Subpart D provides the method for permit writers to
determine when operators qualify for an exemption from the discharge prohibition. For those
operators that quality for the exemption from the discharge prohibition EPA requires that these
operators meet the same stock limitations and discharge limitations for drill cuttings associated
with non-aqueous drilling fluids for operators in Offshore waters (see 40 CFR §435.13 and
§435.15) in order to discharge drill cuttings associated with non-aqueous drilling fluids. This
method remains in 40 CFR 435 and is not included in this document.
The remainder of this section provides a summary of the analytical methods used for Part 435.
1-1
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Table 1-1. Methods by Waste and Pollutant, Subpart A - Offshore Subcategory
Waste source
Produced Water
Water-based drilling
fluids and associated
drill cuttings
Drill cuttings
associated with non-
aqueous drilling
fluids: Discharge
limitations
Regulated Pollutant
Parameter
Oil & grease
SPP Toxicity
Free oil
Diesel oil f
Mercury
Cadmium
Diesel oil f
Free oil
SPP Toxicity
Drilling fluid sediment
toxicity (4-Day Test)
Formation oil
Base fluid retained on
cuttings
Analytical/Test Method
40 CFR Part 136
Drilling Fluids Toxicity Test
EPA Method 1619
Static Sheen Test
EPA Method 16 17
ASTM specification D975-91
40 CFR Part 136
40 CFR Part 136
ASTM specification D975-91
Static Sheen Test
EPA Method 16 17
Drilling Fluids Toxicity Test
EPA Method 1619
Method for Conducting a Sediment Toxicity
Test with Leptocheirus Plumulosus and Non-
Aqueous Drilling Fluids or Synthetic-Based
Drilling Muds (EPA Method 1644)
after using
Procedure for Mixing Base Fluids with
Sediments
EPA Method 1646
Reverse Phase Extraction (RPE) Method for
Detection of Oil Contamination in Non-
Aqueous Drilling Fluids (NAF)
EPA Method 1670
Which can be confirmed by :
Determination of Crude Oil Contamination in
Non-Aqueous Drilling Fluids by Gas
Chromatography/Mass Spectrometry (GC/MS)
EPA Method 1655
Determination of the Amount of Non-Aqueous
Drilling Fluid (NAF) Base Fluid from Drill
Cuttings by a Retort Chamber (Derived from
API Recommended Practice 13B-2)
EPA Method 1674
Cited in 40 CFR
Part
§435.12, §435.13,
§435.14, §435.15
§435.13, §435.15
§435.12, §435.13,
§435.14, §435.15
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.12, §435.14
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
1-2
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Table 1-1. Methods by Waste and Pollutant, Subpart A - Offshore Subcategory
Waste source
Drill cuttings
associated with non-
aqueous drilling
fluids: Stock
limitations (Ci6-C18
internal olefin)
Well treatment,
completion, and
workover fluids
Deck drainage
Domestic waste
Sanitary M10
Sanitary M9IM
Regulated Pollutant
Parameter
Mercury
Cadmium
Polynuclear Aromatic
Hydrocarbons (PAH)
Base fluid sediment
toxicity (10-Day Test)
Biodegradation rate
Oil and grease
Free oil
Free oil
Foam
Floating solids
All other domestic
waste
Total residual chlorine
Floating solids
Analytical/Test Method
40 CFR Part 136
40 CFR Part 136
PAH Content of Oil by HPLC/UV
EPA Method 1654, Revision A
Method for Conducting a Sediment Toxicity
Test with Leptocheirus Plumulosus and Non-
Aqueous Drilling Fluids or Synthetic-Based
Drilling Muds (EPA Method 1644)
after using
Procedure for Mixing Base Fluids with
Sediments
EPA Method 1646
Protocol for the Determination of Degradation
of Non-Aqueous Base Fluids in a Marine
Closed Bottle Biodegradation Test System:
ModifiedlSO 11734:1995
EPA Method 1647
40 CFR Part 136
Static Sheen Test
EPA Method 16 17
Visual sheen
Observation
Observation
See 33 CFR part 15 1{
40 CFR Part 136
Observation
Cited in 40 CFR
Part
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.13, §435.15
§435.12, §435.14
§435.12, §435.13,
§435.14, §435.15
§435.13, §435.15
§435.14, §435.15
§435.14, §435.15
§435.12, §435.14,
§435.15
§435.12, §435.14,
§435.15
§435.12-Off shore BPT
§435.13-OffshoreBAT
§435.14-Offshore BCT
§435.15-Offshore NSPS
•f There is no discharge of diesel oil (see §435.13 and §435.15). Diesel oil refers to the grade of distillate fuel oil, as
specified in ASTM D975-91.
{ This standard references a U.S. Coast Guard regulation: Vessels Carrying Oil, Noxious Liquid Substances,
Garbage, Municipal or Commercial Waste, And Ballast Water (33 CFR part 151).
Note: The Offshore Subcategory bans the discharge of the following waste sources and thus there are no related
analytical or test methods: non-aqueous drilling fluids (NAFs) and produced sand (see §435.12, §435.13, §435.14,
§435.15).
1-3
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Table 1-2. Methods by Waste and Pollutant, Subpart D - Coastal Subcategory
Waste source
Produced Water
Water-based drilling
fluids, drill cuttings,
and dewatering
effluent
Well treatment,
workover, and
completion fluids
Deck drainage
Domestic waste
Sanitary M10
Sanitary M9IM
Regulated Pollutant
Parameter
Oil & grease
SPP Toxicity
Free oil
Diesel oil f
Mercury
Cadmium
Oil and grease
Free oil
Free oil
Foam
Floating solids and
garbage
Total residual chlorine
Floating solids
Analytical/Test Method
40 CFR Part 136
Drilling Fluids Toxicity Test
EPA Method 1619
Static Sheen Test
EPA Method 16 17
ASTM specification D975-91
40 CFR Part 136
40 CFR Part 136
40 CFR Part 136
Static Sheen Test
EPA Method 16 17
Visual sheen
Observation
Observation
40 CFR Part 136
Observation
Cited in 40 CFR
Part
§435.42, §435.43,
§435.44, §435.45
§435.43, §435.45
§435.42, §435.43,
§435.44, §435.45
§435.43, §435.45
§435.43, §435.45
§435.43, §435.45
§435.43, §435.45
§435.42, §435.44
§435.42,§435.43,
§435.44, §435.45
§435.43, §435.45
§435.42, §435.44,
§435.45
§435.42, §435.44,
§435.45
§435.42, §435.44,
§435.45
§435.42 - Coastal BPT §435.44 - Coastal BCT
§435.43 - Coastal BAT §435.45 - Coastal NSPS
f There is no discharge of diesel oil (see §435.43 and §435.45). Diesel oil refers to the grade of distillate fuel oil, as
specified in ASTM D975-91.
Notes:
(1) The Coastal Subcategory bans the discharge of the following waste sources and thus there are no related
analytical or test methods: non-aqueous drilling fluids (NAFs) and produced sand (see §435.42, §435.43, §435.44,
§435.45).
(2) The Coastal Subcategory bans the discharge of drill cuttings associated with non-aqueous drilling fluids unless
there are technical limitations (see §435.43 and §435.45). Appendix 1 of 40 CFR Part 435 Subpart D provides the
method for permit writers to determine when operators qualify for an exemption from this discharge prohibition. For
those operators that quality for the exemption from the discharge prohibition EPA requires that these operators meet
the same stock limitations and discharge limitations for drill cuttings associated with non-aqueous drilling fluids for
operators in Offshore waters (see 40 CFR §435.13 and §435.15) and no discharge of free oil (determined by the
Static Sheen Test, required by §435.42, §435.44) in order to discharge drill cuttings associated with non-aqueous
drilling fluids.
1-4
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1.1 Static Sheen Test (EPA Method 1617)
Scope and Application
This method is to be used as a compliance test for the "no discharge of free oil" requirement for
discharges of drilling fluids, drill cuttings, and well treatment, completion and workover fluids.
"Free oil" refers to any oil contained in a waste stream that when discharged will cause a film or
sheen upon or a discoloration of the surface of the receiving water.
Summary of Method
Samples (15-mL) of drilling fluids or well treatment, completion, and workover fluids, and 15-g
samples (wet weight basis) of drill cuttings or produced sand are introduced into ambient
seawater in a container having an air-to-liquid interface area of 1,000 cm2 (155.5 in2). Samples
are dispersed within the container and observations made no more than one hour later to
ascertain if these materials cause sheen, iridescence, gloss, or increased reflectance on the
surface of the test seawater. The occurrence of any of these visual observations will constitute a
demonstration that the tested material contains "free oil," and therefore results in a prohibition of
its discharge into receiving waters.
1.2 Drilling Fluids Toxicity Test (EPA Method 1619)
Scope and Application
This method is to be used as a compliance test for the suspended particulate phase (SPP) toxicity
requirement for discharges of water-based drilling fluids, associated drill cuttings, and
dewatering effluent and for drill cuttings associated with non-aqueous drilling fluids. The test
may be conducted as a full bioassay to estimate the concentration that is lethal to 50% of the test
organisms (LCso) or as a partial test to determine compliance with 40 CFR Part 435
requirements.
Summary of Method
Samples of drilling fluids or drilling mud from active field systems are diluted with natural or
artificial seawater, mixed, and settled for 1 hour. The suspended particulate phase (SPP) is
decanted. The decanted solution, defined to be 100% SPP, is further diluted with seawater to
obtain test concentrations. Mysidopsis bahia (mysid shrimp) are exposed to the test
concentrations for 96 hours, at which time the living organisms are counted. For the full
bioassay, a range of concentrations is tested and the LCso is calculated. For the partial test, a
negative control, 3% concentration, and positive control are tested. If the number of organisms
killed in the 3% test concentration meets a number specified in the method, the test material
passes the partial toxicity test.
1.3 Procedure for Mixing Base Fluids with Sediments (EPA Method 1646)
This method is used to amend uncontaminated and nontoxic (control) sediments with the base
fluids that are used to formulate synthetic-based drilling fluids and other non-aqueous drilling
fluids. Initially, control sediments are press-sieved through a 2,000 micron mesh sieve to remove
large debris. Then the control sediments are press-sieved through a 500 micron sieve to remove
indigenous organisms that may prey on the test species or otherwise confound test results. The
control sediment is homogenized to limit the effects of settling that may have occurred during
storage. The control sediments are homogenized before density determinations and before the
addition of base fluid. Because base fluids are strongly hydrophobic and do not readily mix with
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sediment, care must be taken to ensure base fluids are thoroughly homogenized within the
sediment. All concentrations are weight-to-weight (mg of base fluid to kg of dry control
sediment).
1.4 Protocol for the Determination of Degradation of Non-Aqueous Base Fluids in a
Marine Closed Bottle Biodegradation Test System: Modified ISO 11734:1995 (EPA
Method 1647)
Scope and Application
EPA promulgated the use of the marine anaerobic biodegradation analytic method because it
most closely modeled the ability of a drilling fluid to biodegrade anaerobically in marine
environments (January 22, 2001; 66 FR 6864). This method determines the anaerobic
degradation potential of mineral oils, paraffin oils and non-aqueous fluids (NAF) in sediments.
These substrates are base fluids for formulating offshore drilling fluids. The test evaluates base
fluid biodegradation rates by monitoring headspace gas production due to microbial degradation
of the test fluid in natural marine sediment.
Subsequent to this promulgation, EPA incorporated additional quality assurance procedures for
the marine anaerobic biodegradation analytic method in the NPDES permit for the Western Gulf
of Mexico ("Final NPDES General Permit for New and Existing Sources and New Dischargers
in the Offshore Subcategory of the Oil and Gas Extraction Category for the Western Portion of
the Outer Continental Shelf of the Gulf of Mexico," GMG290000, Appendix B). The additional
quality assurance instructions in the GMG290000 more clearly describe the sample preparation
and compliance determination steps. Specifically, these additional quality assurance procedures
clarify that users must only use headspace gas to determine compliance with the Part 435
effluent guidelines. EPA noticed its intention to include these additional quality assurance
procedures into Part 435 in the "Guidelines Establishing Test Procedures for the Analysis of
Pollutants Under the Clean Water Act" proposed rulemaking (September 23, 2010; 75 FR
58023). After responding to public comments EPA included these additional quality assurance
procedures into Part 435 with the final rule.
Summary of Method
A mixture of marine/estuarine sediment, test substrate (hydrocarbon or controls), and seawater is
placed into clean 120 ml (150 ml actual volume) Wheaton serum bottles. The test is run using
four replicate serum bottles containing 2,000 mg carbon/kg dry weight concentration of test
substrate in sediment. The anaerobic (redox) condition of the bottles is evaluated using resazurin
dye solution (dye is blue when oxygen is present, reddish in low oxygen conditions and colorless
if oxygen free). Headspace air is removed with a nitrogen sparge before incubation begins. Gas
production and composition are measured approximately every two weeks. Gas production is
measured using a pressure gauge. Barometric pressure is measured at the time of testing to make
necessary volume adjustments. The test period is 275 days. The results of EPA Method 1647 for
the test fluid are compared against the reference fluid to determine whether the test fluid is
eligible for discharge with drill cuttings.
1-6
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1.5 Determination of Crude Oil Contamination in Non-Aqueous Drilling Fluids by Gas
Chromatography/Mass Spectrometry (GC/MS) (EPA Method 1655)
Scope and Application
This method determines crude (formation) oil contamination, or other petroleum oil
contamination, in non-aqueous drilling fluids (NAFs) by comparing the gas
chromatography/mass spectrometry (GC/MS) fingerprint scan and extracted ion scans of the test
sample to that of an uncontaminated sample. It can be used for monitoring oil contamination of
NAFs or monitoring oil contamination of the base fluid used in the NAF formulations.
Summary of Method
Analysis of NAF for crude oil contamination is a step-wise process. The analyst first performs a
qualitative assessment of the presence or absence of crude oil in the sample. If crude oil is
detected during this qualitative assessment, the analyst must perform a quantitative analysis of
the crude oil concentration. A sample of NAF is centrifuged to obtain a solids free supernate.
The test sample is prepared by removing an aliquot of the solids free supernate, spiking it with
internal standard, and analyzing it using GC/MS techniques. The components are separated by
the gas chromatograph and detected by the mass spectrometer.
Qualitative identification of crude oil contamination is performed by comparing the Total Ion
Chromatograph (TIC) scans and Extracted Ion Profile (EIP) scans of test sample to that of
uncontaminated base fluids, and examining the profiles for chromatographic signatures
diagnostic of oil contamination. The presence or absence of crude oil contamination observed in
the full scan profiles and selected extracted ion profiles determines further sample quantitation
and reporting requirements. If crude oil is detected in the qualitative analysis, quantitative
analysis must be performed by calibrating the GC/MS using a designated NAF spiked with
known concentrations of a designated oil.
1.6 Reverse Phase Extraction (RPE) Method for Detection of Oil Contamination in Non-
Aqueous Drilling Fluids (NAF) (EPA Method 1670)
Scope and Application
This method is used for determination of crude or formation oil, or other petroleum oil
contamination, in non-aqueous drilling fluids (NAFs). It is intended as a positive/negative test to
determine a presence of crude oil in NAF prior to discharging drill cuttings from offshore
production platforms. It is for use in the Environmental Protection Agency's (EPA's) survey and
monitoring programs under the Clean Water Act, including monitoring of compliance with the
Gulf of Mexico NPDES General Permit for monitoring of oil contamination in drilling fluids.
The method has been designed to show positive contamination for 5% of representative crude
oils at a concentration of 0.1% in drilling fluid (vol/vol), 50% of representative crude oils at a
concentration of 0.5%, and 95% of representative crude oils at a concentration of 1%.
Summary of Method
An aliquot of drilling fluid is extracted using isopropyl alcohol. The mixture is allowed to settle
and then filtered to separate out residual solids. An aliquot of the filtered extract is charged onto
a reverse phase extraction (RPE) cartridge. The cartridge is eluted with isopropyl alcohol. Crude
oil contaminates are retained on the cartridge and their presence (or absence) is detected based
on observed fluorescence using a black light.
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1.7 Determination of the Amount of Non-Aqueous Drilling Fluid (NAF) Base Fluid from
Drill Cuttings by a Retort Chamber (Derived From API Recommended Practice 13B-
2) (EPA Method 1674)
Scope and Application
This procedure is specifically intended to measure the amount of non-aqueous drilling fluid
(NAF) base fluid from cuttings generated during a drilling operation. This procedure is a retort
test which measures all oily material (NAF base fluid) and water released from a cuttings sample
when heated in a calibrated and properly operating "Retort" instrument.
Summary of Method
A known mass of cuttings is heated in the retort chamber to vaporize the liquids associated with
the sample. The NAF base fluid and water vapors are then condensed, collected, and measured in
a precision graduated receiver.
1.8 PAH Content of Oil by HPLC/UV (EPA Method 1654, Revision A)
Scope and Application
This method is designed to determine the polynuclear aromatic hydrocarbon (PAH) content of
oil by high-performance liquid chromatography (HPLC) with a ultra-violet absorption (UV)
detector. The PAH content is measured and reported as phenanthrene. For oil in drilling muds,
this method is designed to be used in conjunction with the extraction procedure in EPA Method
1662.
Summary of Method
An oil sample is diluted in acetonitrile and a 20-uL aliquot is injected into the HPLC. The
PAHs are partially separated by HPLC and detected with the UV detector. Identification of PAH
(qualitative analysis) is performed by comparing the response of the UV detector to the response
during the retention-time range characteristic of the PAH in diesel oil. PAH is present when a
response occurs during this retention-time range. Quantitative analysis is performed by
calibrating the HPLC with phenanthrene using an external standard technique, and using the
calibration factor to determine the concentration of PAH in the sample.
1.9 Sediment Toxicity Test for NAF and SBM (EPA Method 1644)
Scope and Application
This test method describes the procedures for obtaining data regarding the effects of non-
aqueous drilling fluids (NAFs) or synthetic based drilling muds (SBMs) on the marine
amphipod, Leptocheirusplumulosus. EPA regulates the sediment toxicity in NAFs and SBMs
that are discharged from oil and gas extraction facilities in coastal and offshore waters as an
indicator for toxic pollutants (see 40 CFR 435.13, 435.15, 435.43, and 435.45). EPA established
the use of this test and related limits to encourage the use of less toxic drilling fluids and
additives. The sediment toxicity of the NAF-cuttings at the point of discharge is measured by
this modified sediment toxicity test using a natural sediment and Leptocheirus plumulosus as the
test organism.
EPA promulgated the use of this method with revisions to the Oil and Gas Extraction effluent
guidelines (January 22, 2001; 66 FR 6849). This initial method was consistent with ASTM
Standard Guide E 1367-92 (ASTM 1997). Subsequent to this rulemaking EPA updated this
Ts
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method in the NPDES permit for the Western Gulf of Mexico (GMG290000, Appendix A) to be
consistent with ASTM E 1367-99. Section 10 of this document re-prints the sediment toxicity
test from the NPDES permit for the Western Gulf of Mexico (GMG290000, Appendix A).
Summary of Method
Samples of NAFs and SBMs are exposed to cultured Leptocheirusplumulosus test organisms.
This method relies on standardized bioassay benthic toxicity procedures from ASTM E 1367 and
EPA/600/R-94/0258 along with modified sediment mixing procedures that address the unique
properties of SBM (EPA Method 1646). Of the several species typically used for these tests,
Leptocheirus plumulosus was selected because it demonstrated the range of sensitivity to the
base fluids that were under evaluation and provided both repeatability and discriminatory power.
L. plumulosus live in fine-grained sediments and warm waters similar to the conditions found in
the anticipated receiving environment of the Gulf of Mexico. L. plumulosus is also easy to
culture in the lab and therefore available year-round for testing. The only SBM drill cuttings that
can be discharge are cuttings coated with NAFs and SBM that are as toxic or less toxic, but not
more toxic, than the reference NAF and SBM (Cie-Cig internal olefin or Ci2-Ci4 or Cg ester) as
measured by LCso. This method tests the whole SBM (96-hour exposure test, discharge
limitation) and the NAF (10-day exposure test, stock limitations). This method is referenced in
40 CFR 435.13, 435.15, 435.43, and 435.45.
1.10 Previous Publication of Oil and Gas Extraction Point Source Category Analytic
Methods
EPA previously published the analytical methods for the Oil and Gas Extraction Point Source
Category in the Code of Federal Regulations (CFR). These methods were published as
attachments to Subpart A (Offshore Subcategory). Table 1-3 provides a listing of these methods
as they appear in the CFR and their current EPA method number.
Subsequent to these rulemakings EPA proposed several changes to Part 435, Oil and Gas
Extraction Point Source Category on 23 September 2010 (75 FR 58023). EPA solicited comment
on moving, and in two cases revising, the methods from 40 CFR Part 435, Subpart A (Offshore
Subcategory) to this EPA document, which was included in the rulemaking record for the
proposed rule.1 EPA proposed to organize the analytical methods for the Offshore Subcategory
into this document in order to allow for easier access to the methods for this category. EPA also
proposed to incorporate additional quality assurance procedures in the marine anaerobic
biodegradation analytic method (Appendix 4 to Subpart A of Part 435) and to correct some
erroneous references and omissions in the method for identification of crude oil contamination
(Appendix 5 to Subpart A of Part 435). EPA noticed its intent to include these revisions to these
two methods in the previous version of this document.
EPA received three sets of comments on the above-proposed actions. EPA reviewed and
responded to these comments (see EPA-HQ-OW-2010-0192-0101, EPA-HQ-OW-2010-0192-
0155, EPA-HQ-OW-2010-0192-0127, and the Response to Comments Document, Docket No.
EPA-HQ-OW-2010-0192). In the final rulemaking (2011) EPA retained the full publication of
the Oil and Gas Extraction analytical methods (Appendix 1 through 7 to Subpart A) from 40
1 See "Guidelines Establishing Test Procedures for the Analysis of Pollutants Under the Clean Water Act; Analysis
and Sampling Procedures," PJN 2040-AF09, Docket No. EPA-HQ-OW-2010-0192-0034. Available at:
http://www.epa.gov/regulations.
1-9
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CFR 435, Subpart A, and incorporated these methods by reference to this document. Like any
other changes to an EPA approved method, any changes to the methods in this document will
require a rulemaking.
Table 1-3. EPA Method Numbers for Oil and Gas Extraction Point Source Category
Analytical Methods and Prior CFR References in 40 CFR 435
Analytical/Test Method
Static Sheen Test
Drilling Fluids Toxicity Test
Procedure for Mixing Base Fluids With Sediments
Protocol for the Determination of Degradation of
Non-Aqueous Base Fluids in a Marine Closed Bottle
Biodegradation Test System: Modified ISO
11734:1995
Determination of Crude Oil Contamination in Non-
Aqueous Drilling Fluids by Gas
Chromatography/Mass Spectrometry (GC/MS)
Reverse Phase Extraction (RPE) Method for Detection
of Oil Contamination in Non- Aqueous Drilling Fluids
(NAF)
Determination of the Amount of Non- Aqueous
Drilling Fluid (NAF) Base Fluid from Drill Cuttings
by a Retort Chamber (Derived from API
Recommended Practice 13B-2)
Method for Conducting a Sediment Toxicity Test with
Leptocheirus Plumulosus and Non- Aqueous Drilling
Fluids or Synthetic -Based Drilling Muds
EPA Method
Number
1617
1619
1646
1647
1655
1670
1674
1644
Date First
Promulgated
1993
1993
2001
2001
2001
2001
2001
2001
CFR References
Subpart A, Appendix 1
Subpart A, Appendix 2
Subpart A, Appendix 3
Subpart A, Appendix 4
Subpart A, Appendix 5
Subpart A, Appendix 6
Subpart A, Appendix 7
N/Af
t Note: EPA Method 1644 (sediment toxicity test) was first promulgated for use in 2001 but was incorporated by
reference. Subsequent to this rulemaking EPA updated this method in the NPDES permit for the Western Gulf of
Mexico (GMG290000, Appendix A) to be consistent with ASTM E 1367-99 (ASTM 2000). Section 10 of this
document re-prints the sediment toxicity test from the NPDES permit for the Western Gulf of Mexico
(GMG290000, Appendix A).
2 Federal regulations at 1 CFR 51.7 (c)(l) prohibit the incorporation by reference of material published previously
in the Federal Register. These regulations prevent EPA from deleting these methods from 40 CFR 435.
Ho
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2. STATIC SHEEN TEST (EPA METHOD 1617)
2.1 Scope and Application
This method is to be used as a compliance test for the "no discharge of free oil" requirement for
discharges of drilling fluids, drill cuttings, produced sand, and well treatment, completion and
workover fluids. "Free oil" refers to any oil contained in a waste stream that when discharged
will cause a film or sheen upon or a discoloration of the surface of the receiving water.
2.2 Summary of Method
15-mL samples of drilling fluids or well treatment, completion, and workover fluids, and 15-g
samples (wet weight basis) of drill cuttings or produced sand are introduced into ambient
seawater in a container having an air-to-liquid interface area of 1,000 cm2 (155.5 in2). Samples
are dispersed within the container and observations made no more than one hour later to
ascertain if these materials cause a sheen, iridescence, gloss, or increased reflectance on the
surface of the test seawater. The occurrence of any of these visual observations will constitute a
demonstration that the tested material contains "free oil," and therefore results in a prohibition of
its discharge into receiving waters.
2.3 Interferences
Residual "free oil" adhering to sampling containers, the magnetic stirring bar used to mix the
sample, and the stainless steel spatula used to mix the sample will be the principal sources of
contamination problems. These problems should only occur if improperly washed and cleaned
equipment are used for the test. The use of disposable equipment minimizes the potential for
similar contamination from pipettes and the test container.
2.4 Apparatus, Materials, and Reagents
2.4.1 Apparatus
2.4.1.1 Sampling Containers: 1-liter polyethylene beakers and 1-liter glass
beakers.
2.4.1.2 Graduated cylinder: 100-mL graduated cylinder required only for
operations where predilution of mud discharges is required.
2.4.1.3 Plastic disposable weighing boats.
2.4.1.4 Triple-beam scale.
2.4.1.5 Disposable pipettes: 25-mL disposable pipettes.
2.4.1.6 Magnetic stirrer and stirring bar.
2.4.1.7 Stainless steel spatula.
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2.4.1.8 Test container: Open plastic container whose internal cross-section
parallel to its opening has an area of 1,000 cm2 ±50 cm2 (155.5 ±7.75
in2), and a depth of at least 13 cm (5 inches) and no more than 30 cm (11.8
inches).
2.4.2 Materials and Reagents.
2.4.2.1 Plastic liners for the test container: Oil-free, heavy-duty plastic trash can
liners that do not inhibit the spreading of an oil film. Liners must be of
sufficient size to completely cover the interior surface of the test container.
Permittees must determine an appropriate local source of liners that do not
inhibit the spreading of 0.05 mL of diesel fuel added to the lined test
container under the test conditions and protocol described below.
2.4.2.2 Ambient receiving water.
2.5 Calibration
None currently specified.
2.6 Quality Control Procedures
None currently specified.
2.7 Sample Collection and Handling
2.7.1 Sampling containers must be thoroughly washed with detergent, rinsed a minimum of
three times with fresh water, and allowed to air dry before samples are collected.
2.7.2 Samples of drilling fluid to be tested shall be taken at the shale shaker after cuttings
have been removed. The sample volume should range between 200 mL and 500 mL.
2.7.3 Samples of drill cuttings will be taken from the shale shaker screens with a clean
spatula or similar instrument and placed in a glass beaker. Cuttings samples shall be
collected prior to the addition of any washdown water and should range between
200 g and 500 g.
2.7.4 Samples of produced sand must be obtained from the solids control equipment from
which the discharge occurs on any given day and shall be collected prior to the
addition of any washdown water; samples should range between 200 g and 500 g.
2.7.5 Samples of well treatment, completion, and workover fluids must be obtained from
the holding facility prior to discharge; the sample volume should range between
200 mL and 500 mL.
2.7.6 Samples must be tested no later than 1 hour after collection.
2.7.7 Drilling fluid samples must be mixed in their sampling containers for 5 minutes prior
to the test using a magnetic bar stirrer. If predilution is imposed as a permit condition,
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the sample must be mixed at the same ratio with the same prediluting water as the
discharged muds and stirred for 5 minutes.
2.7.8 Drill cuttings must be stirred and well mixed by hand in their sampling containers
prior to testing, using a stainless steel spatula.
2.8 Procedure
2.8.1 Ambient receiving water must be used as the "receiving water" in the test. The
temperature of the test water shall be as close as practicable to the ambient conditions
in the receiving water, not the room temperature of the observation facility. The test
container must have an air-to-liquid interface area of 1,000 ± 50 cm The surface of
the water should be no more than 1.27 cm (0.5 inch) below the top of the test
container.
2.8.2 Plastic liners shall be used, one per test container, and discarded afterwards. Some
liners may inhibit spreading of added oil; operators shall determine an appropriate
local source of liners that do not inhibit the spreading of the oil film.
2.8.3 A 15-mL sample of drilling fluid or well treatment, completion, and workover fluids
must be introduced by pipette into the test container 1 cm below the water surface.
Pipettes must be filled and discharged with test material prior to the transfer of test
material and its introduction into test containers. The test water/test material mixture
must be stirred using the pipette to distribute the test material homogeneously
throughout the test water. The pipette must be used only once for a test and then
discarded.
2.8.4 Drill cuttings or produced sand should be weighed on plastic weighing boats; 15-g
samples must be transferred by scraping test material into the test water with a
stainless steel spatula. Drill cuttings shall not be prediluted prior to testing. Also,
drilling fluids and cuttings will be tested separately. The weighing boat must be
immersed in the test water and scraped with the spatula to transfer any residual
material to the test container. The drill cuttings or produced sand must be stirred with
the spatula to an even distribution of solids on the bottom of the test container.
2.8.5 Observations must be made no later than 1 hour after the test material is transferred to
the test container. Viewing points above the test container should be made from at
least three sides of the test container, at viewing angles of approximately 60° and 30°
from the horizontal. Illumination of the test container must be representative of
adequate lighting for a working environment to conduct routine laboratory
procedures. It is recommended that the water surface of the test container be observed
under a fluorescent light source such as a dissecting microscope light. The light
source shall be positioned above and directed over the entire surface of the pan.
2.8.6 Detection of a "silvery" or "metallic" sheen or gloss, increased reflectivity, visual
color, iridescence, or an oil slick on the water surface of the test container surface
shall constitute a demonstration of "free oil." These visual observations include
patches, streaks, or sheets of such altered surface characteristics. If the free oil content
of the sample approaches or exceeds 10%, the water surface of the test container may
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lack color, a sheen, or iridescence, due to the increased thickness of the film; thus, the
observation for an oil slick is required. The surface of the test container shall not be
disturbed in any manner that reduces the size of any sheen or slick that may be
present.
If an oil sheen or slick occurs on less than one-half of the surface area after the
sample is introduced to the test container, observations will continue for up to 1 hour.
If the sheen or slick increases in size and covers greater than one-half of the surface
area of the test container during the observation period, the discharge of the material
shall cease. If the sheen or slick does not increase in size to cover greater than one-
half of the test container surface area after one hour of observation, discharge may
continue and additional sampling is not required.
If a sheen or slick occurs on greater than one-half of the surface area of the test
container after the test material is introduced, discharge of the tested material shall
cease. The permittee may retest the material causing the sheen or slick. If subsequent
tests do not result in a sheen or slick covering greater than one-half of the surface area
of the test container, discharge may continue.
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3. DRILLING FLUIDS TOXICITY TEST (EPA METHOD 1619)
3.1 Scope and Application
This method is to be used as a compliance test for the suspended particulate phase (SPP) toxicity
requirement for discharges of water-based drilling fluids, associated drill cuttings, and
dewatering effluent and for drill cuttings associated with non -aqueous drilling fluids. The test
may be conducted as a full bioassay to estimate the concentration that is lethal to 50% of the test
organisms (LCso) or as a partial test to determine compliance with 40 CFR Part 435
requirements.
3.2 Summary of Method
Samples of drilling fluids or drilling mud from active field systems are diluted with natural or
artificial seawater, mixed, and settled for 1 hour. The suspended particulate phase (SPP) is
decanted. The decanted solution, defined to be 100% SPP, is further diluted with seawater to
obtain test concentrations. Mysidopsis bahia (mysid shrimp) are exposed to the test
concentrations for 96 hours, at which time the living organisms are counted. For the full
bioassay, a range of concentrations is tested and the LCso is calculated. For the partial test, a
negative control, 3% concentration, and positive control are tested. If the number of organisms
killed in the 3% test concentration meets a number specified in the method, the test material
passes the partial toxicity test.
3.3 Sample Collection
The collection and preservation methods for drilling fluids (muds) and water samples presented
here are designed to minimize sample contamination and alteration of the physical or chemical
properties of the samples due to freezing, air oxidation, or drying.
3.3.1 Apparatus
3.3.1.1 The following items are required for water and drilling mud sampling and
storage:
a. Acid-rinsed linear-polyethylene bottles or other appropriate
noncontaminating drilling mud sampler.
b. Acid-rinsed linear-polyethylene bottles or other appropriate
noncontaminating water sampler.
c. Acid-rinsed linear-polyethylene bottles or other appropriate
noncontaminated vessels for water and mud samples.
d. Ice chests for preservation and shipping of mud and water samples.
3.3.2 Water Sampling
3.3.2.1 Collection of water samples shall be made with appropriate acid-rinsed
linear-polyethylene bottles or other appropriate non-contaminating water
sampling devices. Special care shall be taken to avoid the introduction of
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contaminants from the sampling devices and containers. Prior to use, the
sampling devices and containers should be thoroughly cleaned with a
detergent solution, rinsed with tap water, soaked in 10% hydrochloric acid
(HC1) for 4 hours, and then thoroughly rinsed with glass-distilled water.
3.3.3 Drilling Mud Sampling
3.3.3.1 Drilling mud formulations to be tested shall be collected from active field
systems. Obtain a well-mixed sample from beneath the shale shaker after
the mud has passed through the screens. Samples shall be stored in
polyethylene containers or in other appropriate uncontaminated vessels.
Prior to sealing the sample containers on the platform, flush as much air
out of the container by filling it with drilling fluid sample, leaving a one
inch space at the top.
3.3.3.2 Mud samples shall be immediately shipped to the testing facility on blue
or wet ice (do not use dry ice) and continuously maintained at 0-4°C until
the time of testing.
3.3.3.3 Bulk mud samples shall be thoroughly mixed in the laboratory using a
1,000 rpm high shear mixer and then subdivided into individual, small
wide-mouthed (e.g., one or two liter) non-contaminating containers for
storage.
3.3.3.4 The drilling muds stored in the laboratory shall have any excess air
removed by flushing the storage containers with nitrogen under pressure
anytime the containers are opened. Moreover, the sample in any container
opened for testing must be thoroughly stirred using a 1,000 rpm high shear
mixer prior to use.
3.3.3.5 Most drilling mud samples may be stored for periods of time longer than 2
weeks prior to toxicity testing provided that proper containers are used and
proper condition are maintained.
3.4 Suspended Particulate Phase Sample Preparation
Mud samples that have been stored under specified conditions in this protocol shall be prepared
for tests within three months after collection. The SPP shall be prepared as detailed below.
3.4.1 Apparatus
3.4.1.1 The following items are required:
a. Magnetic stir plates and bars.
b. Several graduated cylinders, ranging in volume from 10 mL to 1 L
c. Large (15 cm) powder funnels.
d. Several 2-liter graduated cylinders.
O O
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e. Several 2-liter large mouth graduated Erlenmeyer flasks.
3.4.1.2 Prior to use, all glassware shall be thoroughly cleaned. Wash all glassware
with detergent, rinse five times with tap water, rinse once with acetone,
rinse several times with distilled or deionized water, place in a clean 10%
(or stronger) HC1 acid bath for a minimum of 4 hours, rinse five times
with tap water, and then rinse five times with distilled or deionized water.
For test samples containing mineral oil or diesel oil, glassware should be
washed with petroleum ether to assure removal of all residual oil.
Note: If the glassware with nytex cups soaks in the acid solution longer
than 24 hours, then an equally long deionized water soak should be
performed.
3.4.2 Test Seawater Sample Preparation
3.4.2.1 Diluent seawater and exposure seawater samples are prepared by filtration
through a 1.0 micrometer filter prior to analysis.
3.4.2.2 Artificial seawater may be used as long as the seawater has been prepared
by standard methods or ASTM methods, has been properly "seasoned,"
filtered, and has been diluted with distilled water to the same specified
20 ±2 ppt salinity and 20 ±2°C temperature as the "natural" seawater.
3.4.3 Sample Preparation
3.4.3.1 The pH of the mud shall be tested prior to its use. If the pH is less than 9,
if black spots have appeared on the walls of the sample container, or if the
mud sample has a foul odor, that sample shall be discarded. Subsample a
manageable aliquot of mud from the well-mixed original sample. Mix the
mud and filtered test seawater in a volumetric mud-to-water ratio of 1 to 9.
This is best done by the method of volumetric displacement in a 2-L, large
mouth, graduated Erlenmeyer flask. Place 1,000 mL of seawater into the
graduated Erlenmeyer flask. The mud subsample is then carefully added
via a powder funnel to obtain a total volume of 1200 mL. (A 200 mL
volume of the mud will now be in the flask).
The 2-L, large mouth, graduated Erlenmeyer flask is then filled to the
2,000 mL mark with 800 mL of seawater, which produces a slurry with a
final ratio of one volume drilling mud to nine volumes water. If the
volume of SPP required for testing or analysis exceeds 1,500 to 1,600 mL,
the initial volumes should be proportionately increased. Alternatively,
several 2-L drill mud/water slurries may be prepared as outlined above
and combined to provide sufficient SPP.
3.4.3.2 Mix this mud/water slurry with magnetic stirrers for 5 minutes. Measure
the pH and, if necessary, adjust (decrease) the pH of the slurry to within
0.2 units of the seawater by adding 6N HC1 while stirring the slurry. Then,
allow the slurry to settle for 1 hour. Record the amount of HC1 added.
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3.4.3.3 At the end of the settling period, carefully decant (do not siphon) the
Suspended Particulate Phase (SPP) into an appropriate container.
Decanting the SPP is one continuous action. In some cases no clear
interface will be present; that is, there will be no solid phase that has
settled to the bottom. For those samples the entire SPP solution should be
used when preparing test concentrations. However, in those cases when no
clear interface is present, the sample must be remixed for five minutes.
This insures the homogeneity of the mixture prior to the preparation of the
test concentrations. In other cases, there will be samples with two or more
phases, including a solid phase. For those samples, carefully and
continuously decant the supernatant until the solid phase on the bottom of
the flask is reached. The decanted solution is defined to be 100% SPP.
Any other concentration of SPP refers to a percentage of SPP that is
obtained by volumetrically mixing 100% SPP with seawater.
3.4.3.4 SPP samples to be used in toxicity tests shall be mixed for 5 minutes and
must not be preserved or stored.
3.4.3.5 Measure the filterable and unfilterable residue of each SPP prepared for
testing. Measure the dissolved oxygen (DO) and pH of the SPP. If the DO
is less than 4.9 ppm, aerate the SPP to at least 4.9 ppm which is 65% of
saturation. Maximum allowable aeration time is 5 minutes using a generic
commercial air pump and air stone. Neutralize the pH of the SPP to a pH
7.8 ±0.1 using a dilute HC1 solution. If too much acid is added to lower the
pH saturated NaOH may be used to raise the pH to 7.8 ±0.1 units. Record
the amount of acid or NaOH needed to lower/raise to the appropriate pH.
Three repeated DO and pH measurements are needed to insure
homogeneity and stability of the SPP. Preparation of test concentrations
may begin after this step is complete.
3.4.3.6 Add the appropriate volume of 100% SPP to the appropriate volume of
seawater to obtain the desired SPP concentration. The control is seawater
only. Mix all concentrations and the control for 5 minutes by using
magnetic stirrers. Then, the animals shall be randomly selected and placed
in the dishes in order to begin the 96-hour toxicity test.
3.5 Guidance for Performing Suspended Particulate Phase Toxicity Tests Using
Mysidopsis bahia
3.5.1 Apparatus
3.5.1.1 Each definitive test consists of 18 test containers: 3 replicates of a control
and 5 SPP dilutions. Test containers should be Pyrex or equivalent glass.
For definitive tests, 5 SPP dilutions with 3 replicates of at least 500 ml
each are required. Twenty mysids per replicate, 360 per definitive test are
required.
3.5.1.2 A range-finding test consists of at least 6 test containers: 1 replicate of a
control and 5 SPP dilutions. The duration of range-finding tests may be
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less than 96 hours. Ten or more mysids per replicate, at least 60 per range-
finding test, are suggested. Daily water quality measurements and
observations are optional with range-finding tests.
3.5.2 Sample Collection Preservation
3.5.2.1 Drilling muds and water samples are collected and stored as described in
Section 3.3.3.
3.5.3 Species Selection
3.5.3.1 The Suspended Particulate Phase (SPP) tests on drilling muds shall utilize
the test species Mysidopsis bahia. Test animals shall be 3 to 6 days old on
the first day of exposure. Whatever the source of the animals, collection
and handling should be as gentle as possible. Transportation to the
laboratory should be in well-aerated water from the animal culture site at
the temperature and salinity from which they were cultured. Methods for
handling, acclimating, and sizing bioassay organisms given by Borthwick
[1] and Nimmo [2] shall be followed in matters for which no guidance is
given here.
3.5.4 Experimental Conditions
3.5.4.1 Suspended parti culate phase (SPP) tests should be conducted at a salinity
of 20 ±2 ppt. Experimental temperature should be 20 ±2°C. Dissolved
oxygen in the SPP shall be raised to or maintained above 65% of
saturation prior to preparation of the test concentrations. Under these
conditions of temperature and salinity, 65% saturation is a DO of 5.3 ppm.
Beginning at Day 0-before the animals are placed in the test containers
DO, temperature, salinity, and pH shall be measured every 24 hours. DO
should be reported in milligrams per liter.
3.5.4.2 Aeration of test media is required during the entire test with a rate
estimated to be 50-140 cubic centimeters/minute. This air flow to each
test dish may be achieved through polyethylene tubing (0.045-inch inner
diameter and 0.062-inch outer diameter) by a small generic aquarium
pump. The delivery method, surface area of the aeration stone, and flow
characteristics shall be documented. All treatments, including control,
shall be the same.
r\
3.5.4.3 Light intensity shall be 1,200 microwatts/cm using cool white fluorescent
bulbs with a 14-hr light and 10-hr dark cycle. This light/dark cycle shall
also be maintained during the acclimation period and the test.
3.5.5 Experimental Procedure
3.5.5.1 Wash all glassware with detergent, rinse five times with tap water, rinse
once with acetone, rinse several times with distilled or deionized water,
5-5
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place in a clean 10% HC1 acid bath for a minimum of 4 hours, rinse five
times with tap water, and then rinse five times with distilled water.
3.5.5.2 Establish the definitive test concentration based on results of a range
finding test or based on prior experience and knowledge of the mud
system. A minimum of five test concentrations plus a negative and
positive (reference toxicant) control is required for the definitive test. To
estimate the LCso, two concentrations shall be chosen that give (other than
zero and 100%) mortality above and below 50%.
3.5.5.3 Twenty organisms are exposed in each test dish. Nytex® cups shall be
inserted into every test dish prior to adding the animals. These "nylon
mesh screen" nytex holding cups are fabricated by gluing a collar of 363-
micrometer mesh nylon screen to a 15-centimeter wide Petri dish with
silicone sealant. The nylon screen collar is approximately 5 centimeters
high. The animals are then placed into the test concentration within the
confines of the Nytex cups.
3.5.5.4 Individual organisms shall be randomly assigned to treatment. A
randomization procedure is presented in Section 3.7. Make every attempt
to expose animals of approximately equal size. The technique described
by Borthwick [1], or other suitable substitutes, should be used for
transferring specimens. Throughout the test period, mysids shall be fed
daily with approximately 50 Artemia (brine shrimp) nauplii per mysid.
This will reduce stress and decrease cannibalism.
3.5.5.5 Cover the dishes, aerate, and incubate the test containers in an appropriate
test chamber. Positioning of the test containers holding various
concentrations of test solution should be randomized if incubator
arrangement indicates potential position difference. The test medium is not
replaced during the 96-hour test.
3.5.5.6 Observations may be attempted at 4, 6 and 8 hours; they must be
attempted at 0, 24, 48, and 72 hours and must be made at 96 hours.
Attempts at observations refer to placing a test dish on a light table and
visually counting the animals. Do not lift the "nylon mesh screen" cup out
of the test dish to make the observation. No unnecessary handling of the
animals should occur during the 96 hour test period. DO and pH
measurements must also be made at 0, 24, 48, 72, and 96 hours. Take and
replace the test medium necessary for the DO and pH measurements
outside of the nytex cups to minimize stresses on the animals.
3.5.5.7 At the end of 96 hours, all live animals must be counted. Death is the end
point, so the number of living organisms is recorded. Death is determined
by lack of spontaneous movement. All crustaceans molt at regular
intervals, shedding a complete exoskeleton. Care should be taken not to
count an exoskeleton. Dead animals might decompose or be eaten between
observations. Therefore, always count living, not dead animals. If daily
observations are made, remove dead organisms and molted exoskeletons
-------
with a pipette or forceps. Care must be taken not to disturb living
organisms and to minimize the amount of liquid withdrawn.
3.6 Methods for Positive Control Tests (Reference Toxicant)
3.6.1.1 Sodium lauryl sulfate (dodecyl sodium sulfate) is used as a reference
toxicant for the positive control. The chemical used should be
approximately 95% pure. The source, lot number, and percent purity shall
be reported.
3.6.1.2 Test methods are those used for the drilling fluid tests, except that the test
material was prepared by weighing one gram sodium lauryl sulfate on an
analytical balance, adding the chemical to a 100-milliliter volumetric
flask, and bringing the flask to volume with deionized water. After mixing
this stock solution, the test mixtures are prepared by adding 0.1 milliliter
of the stock solution for each part per million desired to one liter of
seawater.
3.6.1.3 The mixtures are stirred briefly, water quality is measured, animals are
added to holding cups, and the test begins. Incubation and monitoring
procedures are the same as those for the drilling fluids.
3.7 Randomization Procedure
3.7.1 Purpose and Procedure
3.7.1.1 The purpose of this procedure is to assure that mysids are impartially
selected and randomly assigned to six test treatments (five drilling fluid or
reference toxicant concentrations and a control) and impartially counted at
the end of the 96-hour test. Thus, each test setup, as specified in the
randomization procedure, consists of 3 replicates of 20 animals for each of
the six treatments, i.e., 360 animals per test. Figure 3-1 is a flow diagram
that depicts the procedure schematically and should be reviewed to
understand the over-all operation. The following tasks shall be performed
in the order listed.
3.7.1.2 Mysids are cultured in the laboratory in appropriate units. If mysids are
purchased, go to Task 3 (Section 3.7.1.3).
3.7.1.3 Remove mysids from culture tanks (6, 5, 4, and 3 days before the test will
begin, i.e., Tuesday, Wednesday, Thursday, and Friday if the test will
begin on Monday) and place them in suitably large maintenance
containers so that they can swim about freely and be fed.
Note: Not every detail (the definition of suitably large containers, for
example) is provided here. Training and experience in aquatic animal
culture and testing will be required to successfully complete these tests.
5-7
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Task
1 Culture Units
2 Maintenance
Containers
3 Test
Population
Containers
4 Separation/
Enumeration
Containers
5 Counting Dish
(repeat tasks
5-7 for A1 and
A2 containers)
Distribution
Containers
7 Test
Containers
Mysids are
collected 3 to
6 days prior
to testing
Figure 3-1. Mysid Randomization Procedure
3.7.1.4 Remove mysids from maintenance containers and place all animals in a
single container. The intent is to have homogeneous test population of
mysids of a known age (3-6 days old).
3.7.1.5 For each toxicity test, assign two suitable containers (500-milliliter (mL)
beakers are recommended) for mysid separation/enumeration. Label each
container (Al, A2, Bl, B2, and Cl, C2, for example, if two drilling fluid
tests and a reference toxicant test are to be set up on one day). The purpose
of this task is to allow the investigator to obtain a close estimate of the
number of animals available for testing and to prevent unnecessary
crowding of the mysids while they are being counted and assigned to test
containers. Transfer the mysids from the large test population container to
the labeled separation and enumeration containers but do not place more
than 200 mysids in a 500-mL beaker. Be impartial in transferring the
-------
mysids; place approximately equal numbers of animals (10-15 mysids is
convenient) in each container in a cyclic manner rather than placing the
maximum number each container at one time.
Note: It is important that the animals not be unduly stressed during this
selection and assignment procedure. Therefore, it will probably be
necessary to place all animals (except the batch immediately being
assigned to test containers) in mesh cups with flowing seawater or in large
volume containers with aeration. The idea is to provide the animals with
near optimal conditions to avoid additional stress.
3.7.1.6 Place the mysids from the two labeled enumeration containers assigned to
a specific test into one or more suitable containers to be used as counting
dishes (2-liter Carolina dishes are suggested). Because of the time required
to separate, count, and assign mysids, two or more people may be
involved in completing this task. If this is done, two or more counting
dishes may be used, but the investigator must make sure that
approximately equal numbers of mysids from each labeled container are
placed in each counting dish.
3.7.1.7 By using a large-bore, smooth-tip glass pipette, select mysids from the
counting dish(es) and place them in the 36 individually numbered
distribution containers (10-ml beakers are suggested). The mysids are
assigned two at a time to the 36 containers by using a randomization
schedule similar to the one presented below. At the end of
selection/assignment round 1, each container will contain two mysids; at
the end of round 2, they will contain four mysids; and so on until each
contains ten mysids.
Table 3-1. Example of a Randomization Schedule
Selection/assignment
round (2 mysids each)
1
2
3
4
5
Place mysid in the numbered distribution containers in the
random order shown
8, 21, 6, 28, 33, 32, 1, 3, 10, 9, 4, 14, 23, 2, 34, 22, 36, 27, 5, 30,
35, 24, 12, 25, 11, 17, 19, 26, 31, 7, 20, 15, 18, 13, 16, 29.
35, 18, 5, 12, 32, 34, 22, 3, 9, 16, 26, 13, 20, 28, 6, 21, 24, 30, 8,
31, 7, 23, 2, 15, 25, 17, 1, 11, 27, 4, 19, 36, 10, 33, 14, 29.
7, 19, 14, 11, 34, 21, 25, 27, 17, 18, 6, 16, 29, 2, 32, 10, 4, 20, 3, 9,
1, 5, 28, 24, 31, 15, 22, 13, 33, 26, 36, 12, 8, 30, 35, 23.
30, 2, 18, 5, 8, 27, 10, 25, 4, 20, 26, 15, 31, 36, 35, 23, 11, 29, 16,
17, 28, 1, 33, 14, 9, 34, 7, 3, 12, 22, 21, 6, 19, 24, 32, 13.
34, 28, 16, 17, 10, 12, 1, 36, 20, 18, 15, 22, 2, 4, 19, 23, 27, 29, 25,
21, 30, 3, 9, 33, 32, 6, 14, 11, 35, 24, 26, 7, 31, 5, 13, 8.
3.7.1.8 Transfer mysids from the 36 distribution containers to 18 labeled test
containers in random order. A label is assigned to each of the three
replicates (A, B, C) of the six test concentrations. Count and record the 96
hour response in an impartial order.
5-9
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3.7.1.9 Repeat Tasks 5-7 (Sections 3.7.1.5 through 3.7.1.7) for each toxicity test.
A new random schedule should be followed in Tasks 6 and 7 for each test.
Note: If a partial toxicity test is conducted, the procedures described above
are appropriate and should be used to prepare the single test concentration
and control, along with the reference toxicant test.
!.7.2 Data Analysis and Interpretation
3.7.2.1 Complete survival data in all test containers at each observation time shall
be presented in tabular form. If greater than 10% mortality occurs in the
controls, all data shall be discarded and the experiment repeated.
Unacceptably high control mortality indicates the presence of important
stresses on the organisms other than the material being tested, such as
injury or disease, stressful physical or chemical conditions in the
containers, or improper handling, acclimation, or feeding. If 10%
mortality or less occurs in the controls, the data may be evaluated and
reported.
3.7.2.2 A definitive, full bioassay conducted according to the EPA protocol is
used to estimate the concentration that is lethal to 50% of the test
organisms that do not die naturally. This toxicity measure is known as the
median lethal concentration, or LCso. The LCso is adjusted for natural
mortality or natural responsiveness. The maximum likelihood estimation
procedure with the adjustments for natural responsiveness as given by DJ.
Finney, in Probit Analysis 3rd edition, 1971, Cambridge University Press,
chapter 7, can be used to obtain the probit model estimate of the LCso and
the 95% fiducial (confidence) limits for the LCso. These estimates are
obtained using the logarithmic transform of the concentration. The
heterogeneity factor (Finney 1971, pages 70-72) is not used. For a test
material to pass the toxicity test, according to the requirements stated in
the offshore oil and gas extraction industry BAT effluent limitations and
NSPS, the LCso, adjusted for natural responsiveness, must be greater than
3% suspended particulate phase (SPP) concentration by volume
unadjusted for the 1 to 9 dilution. Other toxicity test models may be used
to obtain toxicity estimates provided the modeled mathematical expression
for the lethality rate must increase continuously with concentration. The
lethality rate is modeled to increase with concentration to reflect an
assumed increase in toxicity with concentration even though the observed
lethality may not increase uniformly because of the unpredictable animal
response fluctuations.
3.7.2.3 The range finding test is used to establish a reasonable set of test
concentrations in order to run the definitive test. However, if the lethality
rate changes rapidly over a narrow range of concentrations, the range
finding assay may be too coarse to establish an adequate set of test
concentrations for a definitive test.
3-10
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3.7.2.4 The EPA Environmental Research Laboratory in Gulf Breeze, Florida
prepared a Research and Development Report entitled Acute Toxicity of
Eight Drilling Fluids to Mysid Shrimp (Mysidopsis bahia), May 1984
EPA-600/3-84-067. The Gulf Breeze data for drilling fluid number 1 are
displayed in Table 3-2 for purposes of an example of the probit analysis
described above. The SAS Probit Procedure (SAS Institute, Statistical
Analysis System, Gary, North Carolina, 1982) was used to analyze these
data. The 96-hour LCso adjusted for the estimated spontaneous mortality
rate is 3.3% SPP with 95% limits of 3.0 and 3.5% SPP with the 1 to 9
dilution. The estimated spontaneous mortality rate based on all of the data
is 9.6%.
Table 3-2. Listing of Acute Toxicity Test Data (August 1983 to September 1983) With Eight
Generic Drilling Fluids and Mysid Shrimp
[fluid N2=l]
Percent Concentration
0
1
2
3
4
5
Number Exposed
60
60
60
60
60
60
Number Dead (96 hours)
3
11
11
25
48
60
Number Alive (96 hours)
57
49
49
35
12
0
! .7.3 The Partial Toxicity Test for Evaluation of Test Material
3.7.3.1 A partial test conducted according to EPA protocol can be used
economically to demonstrate that a test material passes the toxicity test.
The partial test cannot be used to estimate the LCso adjusted for natural
response.
3.7.3.2 To conduct a partial test, follow the test protocol for preparation of the test
material and organisms. Prepare the control (zero concentration), one test
concentration (3% suspended particulate phase) and the reference toxicant
according to the methods of the full test. A range finding test is not used
for the partial test.
3.7.3.3 Sixty test organisms are used for each test concentration. Find the number
of test organisms killed in the control (zero percent SPP) concentration in
the column labeled XQ of Table 3-3. If the number of organisms in the
control (zero percent SPP) exceeds the table values, then the test is
unacceptable and must be repeated. If the number of organisms killed in
the 3% test concentration is less than or equal to corresponding number in
the column labeled Xi then the test material passes the partial toxicity test.
Otherwise the test material fails the toxicity test.
3.7.3.4 Data shall be reported as percent suspended particulate phase.
3-11
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Table 3-3. Partial Toxicity Test Passing Criteria
Number Killed in Control (X0)
0
1
2
o
3
4
5
6
Number Killed
in Test (Xt)
22
22
23
23
24
24
25
J. 5 References
1. Borthwick, Patrick W. 1978. Methods for acute static toxicity tests with mysid shrimp
(Mysidopsis bahid). Bioassay Procedures for the Ocean Disposal Permit Program, [EPA-
600/9-78-010:] March.
2. Nimmo, D.R., T.L. Hamaker, and C.A. Somers. 1978. Culturing the mysid (Mysidopsis
bahid) in flowing seawater or a static system. Bioassay Procedures for the Ocean
Disposal Permit Program, [EPA-600/9-78-010]: March.
3. American Public Health Association et al. 1980. Standard Methods for the Examination
of Water and Wastewater. Washington, DC, 15th Edition: 90-99.
4. U.S. Environmental Protection Agency, September 1991. Methods for Measuring the
Acute Toxicity of Effluents and Receiving Waters to Freshwater and Marine Organisms.
EPA/600/4-90/027. Washington, DC, 4th Edition.
5. Finney, DJ. Probit Analysis. Cambridge University Press; 1971.
6. U.S. Environmental Protection Agency, May 1984. Acute Toxicity of Eight Drilling
Fluids to Mysid Shrimp (Mysidopsis bahid). EPA-600/3-84-067.
3-12
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4. PROCEDURE FOR MIXING BASE FLUIDS WITH SEDIMENTS (EPA METHOD 1646)
This procedure describes a method for amending uncontaminated and nontoxic (control)
sediments with the base fluids that are used to formulate synthetic-based drilling fluids and other
non-aqueous drilling fluids. Initially, control sediments shall be press-sieved through a 2,000
micron mesh sieve to remove large debris. Then press-sieve the sediment through a 500 micron
sieve to remove indigenous organisms that may prey on the test species or otherwise confound
test results. Homogenize control sediment to limit the effects of settling that may have occurred
during storage. Sediments should be homogenized before density determinations and addition of
base fluid to control sediment. Because base fluids are strongly hydrophobic and do not readily
mix with sediment, care must be taken to ensure base fluids are thoroughly homogenized within
the sediment. All concentrations are weight-to-weight (mg of base fluid to kg of dry control
sediment). Sediment and base fluid mixing shall be accomplished by using the following
method.
4.1 Determining the Wet to Dry Ratio for the Control Sediment
Determine the wet to dry ratio for the control sediment by weighing approximately 10 g
subsamples of the screened and homogenized wet sediment into tared aluminum weigh pans. Dry
sediment at 105°C for 18-24 h. Remove sediment and cool in a desiccator until a constant
weight is achieved. Re-weigh the samples to determine the dry weight. Determine the wet/dry
ratio by dividing the net wet weight by the net dry weight:
Wet Sediment Weight (g) + +
—-——r. . \.f( = Wet to Dry Ratio [4-1 ]
Dry Sediment Weight (g)
4.2 Determining the Density of the Wet Control or Dilution Sediment
Determine the density (g/mL) of the wet control or dilution sediment. This shall be used to
determine total volume of wet sediment needed for the various test treatments.
Mean Wet Sediment Weight (g) „, ^ „ ,. ^_ . , , T, ,._„.,
—:rr „, . g .. ... i /e' = Wet Sediment Density (g/mL) [4-2]
Mean Wet Sediment Volume (mL) J ^° ' L J
4.3 Determining the Amount of Base Fluid Needed
To determine the amount of base fluid needed to obtain a test concentration of 500 mg base fluid
per kg dry sediment, use the following formulas:
Determine the amount of wet sediment required:
Wet Sediment Volume of Sediment Required „ ,. iT, . ,
T->-i^/T\x ^ A *• /- T \ = Sediment Required M ^n
Density (g/mL) per Concentration (mL) „ , ,n V* JJ
J ° ' ^ ^ ' per Cone, (g)
Determine the amount of dry sediment in kilograms (kg) required for each concentration:
4-1
-------
Wet Sediment per
Concentration (g) x 1 kg = Dry Weight Sediment (kg) [4.4]
Mean Wet to Dry Ratio 1,000 g
Finally, determine the amount of base fluid required to spike the control sediment at each
concentration:
Cone. Desired Dry Weight „ r, .,n . ,, , r. ,-.
, ,, , x ? & = Base Fluid Required (mg) [4-5]
(mg/kg) Sediment (kg) n v °'
For spiking test substances other than pure base fluids (e.g., whole mud formulations), determine
the spike amount as follows:
Cone. Desired Dry Weight Test Substance _ Test Substance .-, ,-.
(mL/kg) Sediment (kg) Density (g/mL) Required (g)
4.4 Primary Mixing
For primary mixing, place appropriate amounts of weighed base fluid into stainless mixing
bowls, tare the vessel weight, then add sediment and mix with a high-shear dispersing impeller
for 9 minutes. The concentration of base fluid in sediment from this mix, rather than the nominal
concentration, shall be used in calculating LCso values.
4.5 Testing for Homogeneity of Ease Fluid
Tests for homogeneity of base fluid in sediment are to be performed during the procedure
development phase. Because of difficulty of homogeneously mixing base fluid with sediment, it
is important to demonstrate that the base fluid is evenly mixed with sediment. The sediment shall
be analyzed for total petroleum hydrocarbons (TPH) using EPA Methods 3550A and 8015M,
with samples taken both prior to and after distribution to replicate test containers. Base-fluid
content is measured as TPH. After mixing the sediment, a minimum of three replicate sediment
samples shall be taken prior to distribution into test containers. After the test sediment is
distributed to test containers, an additional three sediment samples shall be taken from three test
containers to ensure proper distribution of base fluid within test containers. Base-fluid content
results shall be reported within 48 hours of mixing. The coefficient of variation (CV) for the
replicate samples must be less than 20%. If base-fluid content results are not within the 20% CV
limit, the test sediment shall be remixed. Tests shall not begin until the CV is determined to be
below the maximum limit of 20%. During the test, a minimum of three replicate containers shall
be sampled to determine base-fluid content during each sampling period.
4.6 Commencing the Sediment Toxicity Test
Mix enough sediment in this way to allow for its use in the preparation of all test concentrations
and as a negative control. When commencing the sediment toxicity test, range-finding tests may
be required to determine the concentrations that produce a toxic effect if these data are otherwise
unavailable. The definitive test shall bracket the LCso, which is the desired endpoint. The results
for the base fluids shall be reported in mg of base fluid per kg of dry sediment.
4-2
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4.7 References
1. American Society for Testing and Materials (ASTM). 1996. Standard Guide for
Collection, Storage, Characterization, and Manipulation of Sediments for Toxicological
Testing. ASTME 1391-94. Annual Book of ASTM Standards, Volume 11.05, pp. SOS-
SIS.
2. Ditsworth, G.R., D.W. Schults and J.K.P. Jones. 1990. Preparation of benthic substrates
for sediment toxicity testing, Environ. Toxicol. Chem. 9:1523-1529.
3. Suedel, B.C., J.H. Rodgers, Jr. and P.A. Clifford. 1993. Bioavailability of fluoranthene in
freshwater sediment toxicity tests. Environ. Toxicol. Chem. 12:155-165.
4. U.S. EPA. 1994. Methods for Assessing the Toxicity of Sediment-associated
Contaminants with Estuarine and Marine Amphipods. EPA/600/R-94/025. Office of
Research and Development, Washington, DC.
5. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category. Federal Register, 66:
6849(22 January 2001).
6. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category: Correction. Federal
Register, 66: 30811 (8 June 2001).
4-3
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5. PROTOCOL FOR THE DETERMINATION OF DEGRADATION OF NON-AQUEOUS BASE
FLUIDS IN A MARINE CLOSED BOTTLE BIODEGRADATION TEST SYSTEM: MODIFIED
ISO 11734:1995 (EPA METHOD 1647)
5.1 Summary of Method
This method determines the anaerobic degradation potential of mineral oils, paraffin oils and
non-aqueous fluids (NAF) in sediments. These substrates are base fluids for formulating offshore
drilling fluids. The test evaluates base fluid biodegradation rates by monitoring gas production
due to microbial degradation of the test fluid in natural marine sediment.
The test procedure places a mixture of marine/estuarine sediment, test substrate (hydrocarbon or
controls) and seawater into clean 120 ml (150 ml actual volume) Wheaton serum bottles. The test
is run using four replicate serum bottles containing 2,000 mg carbon/kg dry weight concentration
of test substrate in sediment. The use of resazurin dye solution (1 ppm) evaluates the anaerobic
(redox) condition of the bottles (dye is blue when oxygen is present, reddish in low oxygen
conditions and colorless if oxygen free). After capping the bottles, a nitrogen sparge removes air
in the headspace before incubation begins. During the incubation period, the sample should be
kept at a constant temperature of 29 ±1°C. Gas production and composition is measured
approximately every two weeks. The samples need to be brought to ambient temperature before
making the measurements. Measure gas production using a pressure gauge. Barometric pressure
is measured at the time of testing to make necessary volume adjustments.
ISO 11734:1995 specifies that total gas is the standard measure of biodegradation. While
modifying this test for evaluating biodegradation of NAFs, methane was also monitored and
found to be an acceptable method of evaluating biodegradation. Section 5.7 contains the
procedures used to follow biodegradation by methane production. Measurement of either total
gas or methane production is permitted. If methane is followed, determine the composition of the
gas by using gas chromatography (GC) analysis at each sampling. At the end of the test when gas
production stops, or at around 275 days, an analysis of sediment for substrate content is possible.
Common methods which have been successfully used for analyzing NAFs from sediments are
listed in Section 5.8.
5.2 System Requirements
This environmental test system has three phases, spiked sediment, overlying seawater, and a gas
headspace. The sediment/test compound mixture is combined with synthetic sea water and
transferred into 120 mL serum bottles. The total volume of sediment/sea water mixture in the
bottles is 75 mL. The volume of the sediment layer will be approximately 50 mL, but the exact
volume of the sediment will depend on sediment characteristics (wetdry ratio and density). The
amount of synthetic sea water will be calculated to bring the total volume in the bottles to 75 mL.
The test systems are maintained at a temperature of 29 ±1°C during incubation. The test systems
are brought to ambient temperatures prior to measuring pressure or gas volume.
5.2.1 Sample Requirements
The concentration of base fluids is at least 2,000 mg carbon test material/kg dry
sediment. Carbon concentration is determined by theoretical composition based on
the chemical formula or by chemical analysis by ASTM D5291-96. Sediments with
-------
positive, intermediate and negative control substances as well as a Cie -Cig internal
olefm type base fluid will be run in conjunction with test materials under the same
conditions. The positive control is ethyl oleate (CAS 111-62-6), the intermediate
control is 1-hexadecene (CAS 629-73-2), and the negative control is squalane (CAS
111-01-3). Controls must be of analytical grade or the highest grade available. Each
test control concentration should be prepared according to the mixing procedure
described in Section 5.3.1.
Product names will be used for examples or clarification in the following text. Any
use of trade or product names in this publication is for descriptive use only, and does
not constitute endorsement by EPA or the authors.
5.2.2 Seawater Requirements
Synthetic seawater at a salinity of 25 ±1 ppt should be used for the test. The synthetic
seawater should be prepared by mixing a commercially available artificial seawater
mix, into high purity distilled or de-ionized water. The seawater should be aerated
and allowed to age for approximately one month prior to use.
5.2.3 Sediment Requirements
The dilution sediment must be from a natural estuarine or marine environment and be
free of the compounds of interest. The collection location, date and time will be
documented and reported. The sediment is prepared by press-sieving through a 2,000-
micron mesh sieve to remove large debris, then press-sieving through a 500-micron
sieve to remove indigenous organisms that may confound test results. The water
content of the sediment should be less than 60% (w/w) or a wet to dry ratio of 2.5.
The sediment should have a minimum organic matter content of 3% (w/w) as
determined by ASTM D2974-07a (Method A and D and calculate organic matter as
in Section 8.3 of method ASTM D2974-07a).
To reduce the osmotic shock to the microorganisms in the sediment the salinity of the
sediment's pore water should be between 20-30 ppt. Sediment should be used for
testing as soon as possible after field collection. If required, sediment can be stored in
the dark at 4°C with 3-6 inches of overlying water in a sealed container for a
maximum period of 2 months prior to use.
5.3 Test Set Up
The test is set up by first mixing the test or control substrates into the sediment inoculum, then
mixing in seawater to make a pourable slurry. The slurry is then poured into serum bottles, which
are then flushed with nitrogen and sealed.
5.3.1 Mixing Procedure
Because base fluids are strongly hydrophobic and do not readily mix with sediments,
care must be taken to ensure base fluids are thoroughly homogenized within the
sediment. All concentrations are weight-to-weight comparisons (mg of base fluid to
5-2
-------
kg of dry control sediment). Sediment and base fluid mixing will be accomplished by
using the following method.
5.3.1.1 Determine the wet to dry weight ratio for the control sediment by
weighing approximately 10 sub-samples of approximately 1 g each of the
screened and homogenized wet sediment into tared aluminum weigh pans.
Dry sediment at 105°C forl8-24 h. Remove the dried sediments and cool
in a desiccator. Repeat the drying, cooling, and weighing cycle until a
constant weight is achieved (within 4% of previous weight). Re-weigh the
samples to determine the dry weight. Calculate the mean wet and dry
weights of the 10 sub samples and determine the wet/dry ratio by dividing
the mean wet weight by the mean dry weight using Equation 5-1. This is
required to determine the weight of wet sediment needed to prepare the
test samples.
Mean Wet Sediment Weight (g) „_ _ _ .
—— ———7. ,„, • u, ) ; = Wet to Dry Ratio [5.1]
Mean Dry Sediment Weight (g) J LJ 1J
5.3.1.2 Determine the density (g/ml) of the wet sediment. This will be used to
determine total volume of wet sediment needed for the various test
treatments. One method is to tare a 5 ml graduated cylinder and add about
5 ml of homogenized sediment. Carefully record the volume then weigh
this volume of sediment. Repeat a total of three times. To determine the
wet sediment density, divide the weight by volume per the following
formula:
Mean Wet Sediment Weight (g) „, 4 „ ,. ^_ . , . , .
—-: ———T. ,. . & / T. = Wet Sediment Density (g/mL) [5.71
Mean Wet Sediment Volume (mL) J v& ' *--> ZJ
5.3.1.3 Determine the amount of base fluid to be spiked into wet sediment in
order to obtain the desired initial base fluid concentration of 2,000 mg
carbon/kg dry weight. An amount of wet sediment that is the equivalent of
30 g of dry sediment will be added to each bottle. A typical procedure is to
prepare enough sediment for 8 serum bottles (3 bottles to be sacrificed at
the start of the test, 4 bottles incubated for headspace analysis, and enough
extra sediment for 2 extra bottles). Extra sediment is needed because some
of the sediment will remain coated onto the mixing bowl and utensils.
Experience with this test may indicate that preparing larger volumes of
spiked sediment is a useful practice. If so, then the following calculations
should be adjusted accordingly.
a. Determine the total weight of dry sediment needed to add 30 g dry
sediment to 8 bottles. If more bottles are used then the calculations
should be modified accordingly. For example:
30 g dry sediment per bottle x 8 = 240 g dry sediment [5-3]
5-3
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b. Determine the weight of base fluid, in terms of carbon, needed to
obtain a final base fluid concentration of 2,000 mg carbon/kg dry
weight. For example:
2,000 mg carbon 240 g .__ ,
5~H A A- ; X i nnn = 48° m§ carbon [5-41
Per kg dry sediment 1,000 ° LJ ^J
c. Convert from mg of carbon to mg of base fluid. This calculation will
depend on the % fraction of carbon present in the molecular structure
of each base fluid. For the control fluids, ethyl oleate is composed of
77.3% carbon, hexadecene is composed of 85.7% carbon, and
squalane is composed of 85.3% carbon. The carbon fraction of each
base fluid should be supplied by the manufacturer or determined
before use. ASTM D5291-96 or equivalent will used to determine
composition of fluid.
To calculate the amount of base fluid to add to the sediment, divide the
amount of carbon (480 mg) by the percent fraction of carbon in the
fluid.
For example, the amount of ethyl oleate added to 240 g dry weight
sediment can be calculated from the following equation:
480 mg carbon , , rr „
(77 3 -H 100) = m§ Y [5"5]
Therefore, add 621 mg of ethyl oleate to 240 g dry weight sediment for
a final concentration of 2,000 mg carbon/kg sediment dry weight.
5.3.1.4 Mix the calculated amount of base fluid with the appropriate weight of wet
sediment.
a. Use the wet:dry ratio to convert from g sediment dry weight to g
sediment wet weight, as follows:
240 g dry sediment xwet:dry ratio = g wet sediment needed [5-6]
b. Weigh the appropriate amount of base fluid (calculated in Section
5.3.1.3c) into stainless mixing bowls, tare the vessel weight, then add
the wet sediment calculated in Equation 5-5, and mix with a high shear
dispersing impeller for 9 minutes.
The sediment is now mixed with synthetic sea water to form a slurry
that will be transferred into the bottles.
5.3.2 Creating Seawater/Sediment Slurry
Given that the total volume of sediment/sea water slurry in each bottle is to be 75 mL,
determine the volume of sea water to add to the wet sediment.
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5.3.2.1 If each bottle is to contain 30 g dry sediment, calculate the weight, and
then the volume, of wet sediment to be added to each bottle.
30 g dry sediment x wetdry ratio = g wet sediment added to each bottle [5-7]
g wet sediment T ,. r
—— . ° T. ,,—-—j. — = mL wet sediment [5-8]
Density (g/mL) or wet sediment
5.3.2.2 Calculate volume of sea water to be added to each bottle.
75 mL total volume - mL wet sediment (from Eq. 8) = mL of sea water [5-9]
5.3.2.3 Determine the ratio of sea water to wet sediment (volume:volume) in each
bottle.
Volume sea water per bottle (Eq. 5-9) _, . ,, ,. r
—77-; T :—f 1T~^—7^—c 0\ = Ratio or sea water:wet sediment [5-10]
Volume sediment water per bottle (Eq. 5-8) L J
5.3.2.4 Convert the wet sediment weight from Equation 5-6 into a volume using
the sediment density.
g wet sediment (Eq. 5-6) density = volume (mL) of sediment [5-11]
5.3.2.5 Determine the amount of sea water to mix with the wet sediment.
mL wet sediment Sea water: sediment ratio T ,, ,. r_ , _..
~ , 11N x ~ , in, = mL sea water to add to wet sediment [5-12]
(Eq. 5-11) (Eq. 5-10) L J
Mix sea water thoroughly with wet sediment to form a sediment/sea water
slurry.
5.3.3 Bottling the Sediment Seawater Slurry
The total volume of sediment/sea water slurry in each bottle is to be 75 mL. Convert
the volume (mL) of sediment/sea water slurry into a weight (g) using the density of
the sediment and the seawater.
5.3.3.1 Determine the weight of sediment to be added to each bottle.
mL sediment (Eq. 5-8) x density of wet sediment (g/mL) = g wet sediment [5-13]
5.3.3.2 Determine the weight of sea water to be added to each bottle.
mL sea water (Eq. 5-9) x density of sea water (1.01 g/mL) = g sea water [5-14]
5.3.3.3 Determine weight of sediment/sea water slurry to be added to each bottle.
g wet sediment (Eq. 5-13) + g sea water (Eq. 5-14) = g sediment/sea water slurry [5-15]
This should provide each bottle with 30 g dry sediment in a total volume of 75 mL.
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5.3.3.4 Putting the sediment:seawater slurry in the serum bottles.
Note: The slurry will need to be constantly stirred to keep the sediment
suspended.
Place a tared serum bottle on a balance and add the appropriate amount of
slurry to the bottle using a funnel. Once the required slurry is in the bottle
remove the funnel, add 2-3 drops (25 uL) of a 1 gram/L resazurin dye
stock solution. Cap the bottle with a butyl rubber stopper (Bellco Glass,
Part #2048- 11800) and crimp with an aluminum seal (Bellco Glass Part
#2048-11020).
Using a plastic tube with a (23 gauge, 1 inch long) needle attached to one
side and a nitrogen source to the other, puncture the serum cap with the
needle. Puncture the serum cap again with a second needle to sparge the
bottle's headspace of residual air for two minutes. The nitrogen should be
flowing at no more than 100 mL/min to encourage gentle displacement of
oxygenated air with nitrogen. Faster nitrogen flow rates would cause
mixing and complete oxygen removal would take much longer. Remove
the nitrogen needle first to avoid any initial pressure problems. The second
(vent) needle should be removed within 30 seconds of removing the
nitrogen needle.
Triplicate blank test systems are prepared, with similar quantities of
sediment and seawater without any base fluid. Incubate in the dark at a
constant temperature of 29 ±1°C.
Record the test temperature. The test duration is dependent on base fluid
performance, but at a maximum should be no more than 275 days. Stop
the test after all base fluids have achieved a plateau of gas production. At
termination, base fluid concentrations can be verified in the terminated
samples by extraction and GC analysis according to Section 5.8.
5.4 Concentration Verification Chemical Analyses
Because of the difficulty of homogeneously mixing base fluid with sediment, it is important to
demonstrate that the base fluid is evenly mixed within the sediment sea water slurry that was
added to each bottle. Of the seven serum bottles set up for each test or control condition, three
are randomly selected for concentration verification analyses. These should be immediately
placed at 4°C and a sample of sediment from each bottle should be analyzed for base fluid
content as soon as possible. The coefficient of variation (CV) for the replicate samples must be
less than 20%. The results should show recovery of at least 70% of the spiked base fluid. Use an
appropriate analytical procedure described in Section 5.8 to perform the extractions and analyses.
If any set of sediments fail the criteria for concentration verification, then the corrective action
for that set of sediments is also outlined in Section 5.8.
The nominal concentrations and the measured concentrations from the three bottles selected for
concentration verification should be reported for the initial test concentrations. The coefficient of
variation (CV) for the replicate samples must be less than 20%. If base fluid content results are
-------
not within the 20% CV limit, the test must be stopped and restarted with adequately mixed
sediment.
5. 5 Gas Monitoring Procedures
Biodegradation is measured by total gas as specified in ISO 11734:1995. Methane
production can also
be tracked and is described in Section 5.7.
5.5.1 Total Gas Monitoring Procedures
Bottles should be brought to room temperature before readings are taken. The bottles
are observed to confirm that the resazurin has not oxidized to pink or blue. Total gas
production in the culture bottles should be measured using a pressure transducer (one
source is Biotech International). The pressure readings from test and control cultures
are evaluated against a calibration curve created by analyzing the pressure created by
known additions of gas to bottles established identically to the culture bottles. Bottles
used for the standard curve contain 75 mL of water, and are sealed with the same
rubber septa and crimp cap seals used for the bottles containing sediment. After the
bottles used in the standard curve have been sealed, a syringe needle inserted through
the septa is used to equilibrate the pressure inside the bottles to the outside
atmosphere. The syringe needle is removed and known volumes of air are injected
into the headspace of the bottles. Pressure readings provide a standard curve relating
the volume of gas injected into the bottles and headspace pressure. No less than three
points may be used to generate the standard curve. A typical standard curve may use
0, 1,5, 10, 20 and 40 ml of gas added to the standard curve bottles.
The room temperature and barometric pressure (to two digits) should be recorded at
the time of sampling. One option for the barometer is Fisher Part #02-400 or 02-401 .
Gas production by the sediment is expressed in terms of the volume (mL) of gas at
standard temperature (0°C = 273°K) and pressure (1 atm = 30 inches of Hg) using Eq.
5-16.
Pi x Vi x T?
T.«pa -
Where:
V2 = Volume of gas production at standard temperature and pressure
PI = Barometric pressure on day of sampling (inches of Hg)
Vi = Volume of gas measured on day of sampling (mL)
T2 = Standard temperature = 273 °K
TI = Temperature on day of sampling (°C + 273 = °K)
?2 = Standard pressure = 30 inches Hg
An estimate can be made of the total volume of anaerobic gas that will be produced in
the bottles. The gas production measured for each base fluid can be expressed as a
percent of predicted total anaerobic gas production.
5-7
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5.5.1.1 Calculate the total amount of carbon in the form of the base fluid present
in each bottle.
Each bottle is to contain 30 g dry weight sediment. The base fluid
concentration is 2,000 mg carbon/kg dry weight sediment. Therefore:
2,000 mg carbon/kg sediment x (30 g ^ 1,000) = 60 mg carbon per bottle [5-1]
5.5.1.2 Theory states that anaerobic microorganisms will convert 1 mole of
carbon substrate into 1 mole of total anaerobic gas production.
Calculate the number of moles of carbon in each bottle.
The molecular weight of carbon is 12 (i.e., 1 mole of carbon = 12 g).
Therefore, the number of moles of carbon in each bottle can be calculated.
60 mg carbon per bottle/1,000
= TTr-T—; = 0.005 moles carbon [5-21
12 g/mole L J
5.5.1.3 Calculate the predicted volume of anaerobic gas.
One mole of gas equals 22.4 L (at standard temperature and pressure),
therefore,
0.005 moles x 22.4 L = 0.112L (or 112 mL total gas production) [5-3]
5.5.2 Gas Venting
If the pressure in the serum bottle is too great for the pressure transducer or syringe,
some of the excess gas must be wasted. The best method to do this is to vent the
excess gas right after measurement. To do this, remove the barrel from a 10-mL
syringe and fill it 1/3 full with water. This is then inserted into the bottle through the
stopper using a small diameter (high gauge) needle. The excess pressure is allowed to
vent through the water until the bubbles stop. This allows equalization of the pressure
inside the bottle to atmospheric without introducing oxygen. The amount of gas
vented (which is equal to the volume determined that day) must be kept track of each
time the bottles are vented. A simple way to do this in a spreadsheet format is to have
a separate column in which cumulative vented gas is tabulated. Each time the volume
of gas in the cultures is analyzed, the total gas produced is equal to the gas in the
culture at that time plus the total of the vented gas.
To keep track of the methane lost in the venting procedure, multiply the amount of
gas vented each time by the corrected % methane determined on that day. The answer
gives the volume of methane wasted. This must be added into the cumulative totals
similarly to the total gas additions.
5.6 Test Acceptability and Interpretation
5.6.1 Test Acceptability
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At day 275 or when gas production has plateaued, whichever is first, the controls are
evaluated to confirm that the test has been performed appropriately. In order for this
modification of the closed bottle biodegradation test to be considered acceptable, all
the controls must meet the biodegradation levels indicated in Table 5-1. The
intermediate control hexadecene must produce at least 30% of the theoretical gas
production. This level may be reexamined after two years and more data has been
generated.
Table 5-1. Test Acceptability Criteria
Concentration
2,000 mg carbon/kg
Percent Biodegradability as a Function of Gas Measurement
Positive control
>60% theoretical
Squalane negative control
<5% theoretical
Hexadecene intermediate control
>30% theoretical
5.6.2 Interpretation
In order for a fluid to pass the closed bottle test, the biodegradation of the base fluid
as indicated by the total amount of total gas (or methane) generated once gas
production has plateaued (or at the end of 275 days, whichever is first) must be
greater than or equal to the volume of gas (or methane) produced by the reference
standard (internal elefin or ester).
The method for evaluating the data to determine whether a fluid has passed the
biodegradation test must use the equations:
% Theoretical gas production of reference fluid
% Theoretical gas production of NAF ~ L - J
Where:
NAF = Stock base fluid being tested for compliance
Reference fluid = Cie -Cig internal olefm or Cu -Ci4 or Cg ester reference
fluid
5.7 Methane Measurement
5.7.1 Methane Monitoring Procedures
The use of total gas production alone may result in an underestimation of the actual
metabolism occurring since CC>2 is slightly soluble in water. An acceptable alternative
method is to monitor methane production and total gas production. This is easily done
using GC analysis. A direct injection of headspace gases can be made into a GC using
almost any packed or capillary column with an FID detector. Unless volatile fuels or
solvents are present in the test material or the inocula, the only component of the
headspace gas that can be detected using an FID detector is methane. The percent
methane in the headspace gas is determined by comparing the response of the sample
injections to the response from injections of known percent methane standards. The
percent methane is corrected for water vapor saturation using Eq. 5-21 and then
converted to a volume of dry methane using Eq. 5-22.
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Corrected % CH4 = %CH4
_ D x 22.4 L/mol „
18g/mol x 1,000 "
Where:
D = The density of water vapor at saturation (g/m3, can be found in
CRC Handbook of Chemistry and Physics] for the temperature
of sampling.
ml) = (S + V) x -- x __ x __ [5.22]
Where:
VcH4 = The volume of methane in the bottle
S = Volume of excess gas production (measured with a pressure
transducer)
V = Volume of the headspace in the culture bottle (total volume -liquid
phase)
P = Barometric pressure (mm Hg, measured with barometer)
T = Temperature (°C)
Pw = Vapor pressure of water at T (mm Hg, can be found in CRC
Handbook of Chemistry and Physics)
CH4 = % methane in headspace gas (after correction for water vapor)
The total volume of serum bottles sold as 125 mL bottles (Wheaton) is 154.8 mL.
The volumes of methane produced are then compared to the volumes of methane in
the controls to determine if a significant inhibition of methane production or a
significant increase of methane production has been observed. Effective statistical
analyses are important, as variability in the results is common due to the
heterogeneity of the inoculum's source. It is also common to observe that the timing
of the initiation of culture activity is not equal in all of the cultures. Expect a great
variability over the period when the cultures are active, some replicates will start
sooner than others, but all of the replicates should eventually reach similar levels of
base fluid degradation and methane production.
5.7.2 Expected Methane Production Calculations
The amount of methane expected can be calculated using the equation of Symons and
Buswell (Eq. 5-23). In the case of complete mineralization, all of the carbon will
appear as wither CC>2 or CH/j, thus the total moles of gas produced will be equal to the
total moles of carbon in the parent molecule. The use of the Buswell equation allows
you to calculate the effects the redox potential will have on the distribution of the
products in methanogenic cultures. More reduced electron donors will allow the
production of more methane, while more oxidized electron donors will cause a
production of more carbon dioxide.
5-10
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CnHaObNcSd + (n-a/4 -b/2 + 7c/4 + d/2) H2O -> (n/2 -a/8+b/4-5c/8 + d/4) CO2 +
(n/2 +a/8 -b/4 -3c/8-d/4) CH4 + cNH4HCO3 + dH2S [5-23]
An example calculation of the expected methane volume in a culture fed 2,000 mg/kg
hexadecene is as follows. The application of Symons and Buswell's equation reveals
that hexadecene (Cielfe) will yield 4 moles of CO2 and 12 moles of CH4. Assuming
30 g of dry sediment are added to the bottles with 2,334 mg hexadecene/kg dry
sediment (i.e., equivalent to 2,000 mg carbon/kg dry sediment) the calculation is as
follows.
1,000 1 mole 23 g
12moleCH4 22.4 L ml hexadecene hexadecene 0.03kg _ . , ,, r^ 0/n
—x x x x x s—= 84 (ml} P-241
mole mole CH4 L 224.4 g kg dry soil culture v '
hexadecene hexadecene
By subtracting the average amount of methane in control bottles from the test bottles
and then dividing by the expected volume an evaluation of the completion of the
process may be conducted.
5.8 Concentration Verification Analysis
The Concentration Verification analysis is required at the beginning of the test to ensure
homogeneity and confirm that the required amount of fluid was delivered to the sediments at the
start of the test.
5.8.1 Three samples per fluid need to be analyzed and achieve <20% Coefficient of
Variability and an average of >70% to <120% of fluid delivered to sediment.
5.8.2 If a third party performs the analysis, then the laboratory should be capable of
delivering the homogeneity data within seven days, in order to identify any samples
that do not meet the homogeneity requirement as quickly as possible.
5.8.3 If one sediment/fluid set, out a multiple set batch of samples, fails these criteria, then
that one set of samples must be discarded and a fresh set of spiked sediment prepared,
started, and analyzed to ensure homogeneity. The same stock sediment is used to
prepare the replacement set(s). The remaining sets do not need to be re-mixed or
restarted.
5.8.4 The re-mixed set(s) will need to be run the additional days as appropriate to ensure
that the total number of days is the same for all sets of bottles, even though the
specific days are not aligned.
5.8.5 Re-mixing of bottle sets can be performed multiple times as a result of a failure of the
analytical criteria, until the holding time for the stock sediment has expired (60 days).
If the problem set(s) has not fallen within the acceptable analytical criteria by then, it
must not be part of the batch of bottles run. If the problem batch is one of the
controls, and those controls were not successfully prepared when the sediment
holding time expired, then the entire test must be restarted.
5-11
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5.9 Program Quality Assurance and Quality Control
5.9.1 Calibration
5.9.1.1 All equipment / instrumentation will be calibrated in accordance with the
test method or the manufacturer's instructions and may be scheduled or
triggered.
5.9.1.2 Where possible, standards used in calibration will be traceable to a
nationally recognized standard (e.g., certified standard by NIST).
5.9.1.3 All calibration activities will be documented and the records retained.
5.9.1.4 The source, lot, batch number, and expiration date of all reagents used
with be documented and retained.
5.9.2 Maintenance
5.9.2.1 All equipment / instrumentation will be maintained in accordance with the
test method or the manufacturer's instructions and may be scheduled or
triggered.
5.9.2.2 All maintenance activities will be documented and the records retained.
5.9.3 Data Management and Handling
5.9.3.1 All primary (raw) data will be correct, complete, without selective
reporting, and will be maintained.
5.9.3.2 Hand-written data will be recorded in lab notebooks or electronically at
the time of observation.
5.9.3.3 All hand-written records will be legible and amenable to reproduction by
electrostatic copiers.
5.9.3.4 All changes to data or other records will be made by:
a. Using a single line to mark-through the erroneous entry (maintaining
original data legibility)
b. Write the revision
c. Initial, date, and provide revision code (see attached or laboratory's
equivalent)
5.9.3.5 All data entry, transcriptions, and calculations will be verified by a
qualified person.
a. Verification will be documented by initials of verifier and date
5-12
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5.9.3.6 Procedures will be in place to address data management procedures used
(at minimum):
a. Significant figures
b. Rounding practices
c. Identification of outliers in data series
d. Required statistics
5.9.4 Document Control
5.9.4.1 All technical procedures, methods, work instructions, standard operating
procedures must be documented and approved by laboratory management
prior to the implementation.
5.9.4.2 All primary data will be maintained by the contractor for a minimum of
five (5) years.
5.9.5 Personnel and Training
5.9.5.1 Only qualified personnel shall perform laboratory activities.
5.9.5.2 Records of staff training and experience will be available. This will
include initial and refresher training (as appropriate).
5.9.6 Test Performance
5.9.6.1 All testing will be done in accordance with the specified test methods.
5.9.6.2 Receipt, arrival condition, storage conditions, dispersal, and accountability
of the test article will be documented and maintained.
5.9.6.3 Receipt or production, arrival or initial condition, storage conditions,
dispersal, and accountability of the test matrix (e.g., sediment or artificial
seawater) will be documented and maintained.
5.9.6.4 Source, receipt, arrival condition, storage conditions, dispersal, and
accountability of the test organisms (including inoculum) will be
documented and maintained.
5.9.6.5 Actual concentrations administered at each treatment level will be verified
by appropriate methodologies.
5.9.6.6 Any data originating at a different laboratory will be identified and the
laboratory fully referenced in the final report.
5.9.7 The following references identify analytical methods that have historically been
successful for achieving the analytical quality criteria.
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5.9.7.1 Continental Shelf Associates report 1998. Joint EPA/Industry screening
survey to assess the deposition of drill cuttings and associated synthetic
based mud on the seabed of the Louisiana continental shelf, Gulf of
Mexico. Analysis by Charlie Henry report Number IES/RCAT97-36 GC-
FID and GC/MS.
5.9.7.2 EPA Method 3550 for extraction with EPA Method 8015 for GC-FID.
EPA Method 3550C, Revision 3, February 2007. Ultrasonic Extraction.
EPA Method 8015C, Revision 3, February 2007. Nonhalogenated
Organics by Gas Chromatography.
5.9.7.3 Chandler, I.E., S.P. Rabke, and A.J.J. Leuterman. 1999. Predicting the
Potential Impact of Synthetic-Based Muds with the Use of Biodegradation
Studies. Society of Petroleum Engineers SPE 52742.
5.9.7.4 Chandler, I.E., B. Lee, S.P. Rabke, J.M. Geliff, R. Stauffer, and J. Hein.
2000. Modification of a Standardized Anaerobic Biodegradation Test to
Discriminate Performance of Various Non-Aqueous Base Fluids. Society
of Petroleum Engineers SPE 61203.
5.9.7.5 Munro P.O., B. Croce, C.F. Moffet, N.A Brown, A.D. Mclntosh, SJ.
Hird, andR.M. Stagg. 1998. Solid-phase test for comparison of
degradation rates of synthetic mud base fluids used in the off-shore
drilling industry. Environ. Toxicol Chem. 17:1951-1959.
5.9.7.6 Webster, L, P.R. Mackie, SJ. Hird, P.O. Munro, N.A. Brown, and C.F.
Moffat. 1997. Development of Analytical Methods for the Determination
of Synthetic Mud Base Fluids in Marine Sediments. Analyst 122:1485-
1490.
5-14
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6. DETERMINATION OF CRUDE OIL CONTAMINATION IN NON-AQUEOUS DRILLING FLUIDS
BY GAS CHROMATOGRAPHY/MASS SPECTROMETRY (GC/MS) (EPA METHOD 1655)
6.1 Scope and Application
6.1.1 This method determines crude (formation) oil contamination, or other petroleum oil
contamination, in non-aqueous drilling fluids (NAFs) by comparing the gas
chromatography/mass spectrometry (GC/MS) fingerprint scan and extracted ion scans
of the test sample to that of an uncontaminated sample.
6.1.2 This method can be used for monitoring oil contamination of NAFs or monitoring oil
contamination of the base fluid used in the NAF formulations.
6.1.3 Any modification of this method beyond those expressly permitted shall be
considered as a major modification subject to application and approval of alternative
test procedures under 40 CFR 136.4 and 136.5.
6.1.4 The gas chromatography/mass spectrometry portions of this method are restricted to
use by, or under the supervision of analysts experienced in the use of GC/MS and in
the interpretation of gas chromatograms and extracted ion scans. Each laboratory that
uses this method must generate acceptable results using the procedures described in
Sections 6.7, 6.9.2, and 6.12.
6.2 Summary of Method
6.2.1 Analysis of NAF for crude oil contamination is a step-wise process. The analyst first
performs a qualitative assessment of the presence or absence of crude oil in the
sample. If crude oil is detected during this qualitative assessment, the analyst must
perform a quantitative analysis of the crude oil concentration.
6.2.2 A sample of NAF is centrifuged to obtain a solids free supernate.
6.2.3 The test sample is prepared by removing an aliquot of the solids free supernate,
spiking it with internal standard, and analyzing it using GC/MS techniques. The
components are separated by the gas chromatograph and detected by the mass
spectrometer.
6.2.4 Qualitative identification of crude oil contamination is performed by comparing the
Total Ion Chromatograph (TIC) scans and Extracted Ion Profile (EIP) scans of test
sample to that of uncontaminated base fluids, and examining the profiles for
chromatographic signatures diagnostic of oil contamination.
6.2.5 The presence or absence of crude oil contamination observed in the full scan profiles
and selected extracted ion profiles determines further sample quantitation and
reporting requirements.
6.2.6 If crude oil is detected in the qualitative analysis, quantitative analysis must be
performed by calibrating the GC/MS using a designated NAF spiked with known
concentrations of a designated oil.
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6.2.7 Quality is assured through reproducible calibration and testing of GC/MS system and
through analysis of quality control samples.
6.3 Definitions
6.3.1 A NAF is one in which the continuous phase is a water immiscible fluid such as an
oleaginous material (e.g., mineral oil, enhance mineral oil, paraffinic oil, or synthetic
material such as olefms and vegetable esters).
6.3.2 TIC—Total Ion Chromatograph.
6.3.3 EIP—Extracted Ion Profile.
6.3.4 TCB—1,3,5-trichlorobenzene is used as the internal standard in this method.
6.3.5 SPTM—System Performance Test Mix standards are used to establish retention times
and monitor detection levels.
6.4 Interferences and Limitations
6.4.1 Solvents, reagents, glassware, and other sample processing hardware may yield
artifacts and/or elevated baselines causing misinterpretation of chromatograms.
6.4.2 All Materials used in the analysis shall be demonstrated to be free from interferences
by running method blanks. Specific selection of reagents and purification of solvents
by distillation in all-glass systems may be required.
6.4.3 Glassware shall be cleaned by rinsing with solvent and baking at 400°C for a
minimum of 1 hour.
6.4.4 Interferences may vary from source to source, depending on the diversity of the
samples being tested.
6.4.5 Variations in and additions of base fluids and/or drilling fluid additives (emulsifiers,
dispersants, fluid loss control agents, etc.) might also cause interferences and
misinterpretation of chromatograms.
6.4.6 Difference in light crude oils, medium crude oils, and heavy crude oils will result in
different responses and thus different interpretation of scans and calculated
percentages.
6.5 Safety
6.5.1 The toxicity or carcinogenicity of each reagent used in this method has not been
precisely determined; however each chemical shall be treated as a potential health
hazard. Exposure to these chemicals should be reduced to the lowest possible level.
6.5.2 Unknown samples may contain high concentration of volatile toxic compounds.
Sample containers should be opened in a hood and handled with gloves to prevent
-------
exposure. In addition, all sample preparation should be conducted in a fume hood to
limit the potential exposure to harmful contaminates.
6.5.3 This method does not address all safety issues associated with its use. The laboratory
is responsible for maintaining a safe work environment and a current awareness file
of OSHA regulations regarding the safe handling of the chemicals specified in this
method. A reference file of material safety data sheets (MSDSs) shall be available to
all personnel involved in these analyses. Additional references to laboratory safety
can be found in Section 6.16, References 1 through 3.
6.5.4 NAF base fluids may cause skin irritation, protective gloves are recommended while
handling these samples.
6.6 Apparatus and Materials
Note: Brand names, suppliers, and part numbers are for illustrative purposes only. No
endorsement is implied. Equivalent performance may be achieved using apparatus and materials
other than those specified here, but demonstration of equivalent performance meeting the
requirements of this method is the responsibility of the laboratory.
6.6.1 Equipment for glassware cleaning.
6.6.1.1 Laboratory sink with overhead fume hood.
6.6.1.2 Kiln—Capable of reaching 450°C within 2 hours and holding 450°C
within ±10°C, with temperature controller and safety switch (Cress
Manufacturing Co., Santa Fe Springs, CA B31H or X3 ITS or equivalent).
6.6.2 Equipment for sample preparation.
6.6.2.1 Laboratory fume hood.
6.6.2.2 Analytical balance—Capable of weighing 0.1 mg.
6.6.2.3 Glassware.
a. Disposable pipettes—Pasteur, 150 mm long by 5 mm ID (Fisher
Scientific 13-678-6A, or equivalent) baked at 400°C for a minimum
of 1 hour.
b. Glass volumetric pipettes or gas tight syringes—1.0-mL ±1% and 0.5-
mL±l%.
c. Volumetric flasks—Glass, class A, 10-mL, 50-mL and 100-mL.
d. Sample vials—Glass, 1- to 3-mL (baked at 400°C for a minimum of 1
hour) with PTFE-lined screw or crimp cap.
e. Centrifuge and centrifuge tubes—Centrifuge capable of 10,000 rpm, or
better, (International Equipment Co., IEC Centra MP4 or equivalent)
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and 50-mL centrifuge tubes (Nalgene, Ultratube, Thin Wall 25x89
mm, #3410-2539).
6.6.3 Gas Chromatograph/Mass Spectrometer (GC/MS):
6.6.3.1 Gas Chromatograph—An analytical system complete with a temperature-
programmable gas chromatograph suitable for split/splitless injection and
all required accessories, including syringes, analytical columns, and gases.
a. Column—30 m (or 60 m) x 0.32 mm ID (or 0.25 mm ID) 1 |im film
thickness (or 0.25 jim film thickness) silicone-coated fused-silica
capillary column (J&W Scientific DB-5 or equivalent).
6.6.3.2 Mass Spectrometer—Capable of scanning from 35 to 600 amu every 1 sec
or less, using 70 volts (nominal) electron energy in the electron impact
ionization mode (Hewlett Packard 5970MS or comparable).
6.6.3.3 GC/MS interface—the interface is a capillary-direct interface from the GC
to the MS.
6.6.3.4 Data system—A computer system must be interfaced to the mass
spectrometer. The system must allow the continuous acquisition and
storage on machine-readable media of all mass spectra obtained
throughout the duration of the chromatographic program. The computer
must have software that can search any GC/MS data file for ions of a
specific mass and that can plot such ion abundance versus retention time
or scan number. This type of plot is defined as an Extracted Ion Current
Profile (EIP). Software must also be available that allows integrating the
abundance in any total ion chromatogram (TIC) or EIP between specified
retention time or scan-number limits. It is advisable that the most recent
version of the EPA/NIST Mass Spectral Library be available.
6.7 Reagents and Standards
6.7.1 Methylene chloride—Pesticide grade or equivalent. Use when necessary for sample
dilution.
6.7.2 Standards—Prepare from pure individual standard materials or purchase as certified
solutions. If compound purity is 96% or greater, the weight may be used without
correction to compute the concentration of the standard.
6.7.2.1 Crude Oil Reference—Obtain a sample of a crude oil with a known API
gravity. This oil shall be used in the calibration procedures.
6.7.2.2 Synthetic Base Fluid—Obtain a sample of clean internal olefm (IO) Lab
drilling fluid (as sent from the supplier—has not been circulated
downhole). This drilling fluid shall be used in the calibration procedures.
6.7.2.3 Internal standard—Prepare a 0.01 g/mL solution of 1,3,5-trichlorobenzene
(TCB). Dissolve 1.0 g of TCB in methylene chloride and dilute to volume
-------
in a 100-mL volumetric flask. Stopper, vortex, and transfer the solution to
a 150-mL bottle with PTFE-lined cap. Label appropriately, and store at
-5°C to 20°C. Mark the level of the meniscus on the bottle to detect
solvent loss.
6.7.2.4 GC/MS system performance test mix (SPTM) standards—The SPTM
standards shall contain octane, decane, dodecane, tetradecane, tetradecene,
toluene, ethylbenzene, 1,2,4-trimethylbenzene, 1-methylnaphthalene and
1,3-dimethylnaphthalene. These compounds can be purchased individually
or obtained as a mixture (i.e., Supelco, Catalog No. 4-7300). Prepare a
high concentration of the SPTM standard at 62.5 mg/mL in methylene
chloride. Prepare a medium concentration SPTM standard at 1.25 mg/mL
by transferring 1.0 mL of the 62.5 mg/mL solution into a 50 mL
volumetric flask and diluting to the mark with methylene chloride. Finally,
prepare a low concentration SPTM standard at 0.125 mg/mL by
transferring 1.0 mL of the 1.25 mg/mL solution into a 10-mL volumetric
flask and diluting to the mark with methylene chloride.
6.7.2.5 Crude oil/drilling fluid calibration standards—Prepare a 4-point crude
oil/drilling fluid calibration at concentrations of 0% (no spike—clean
drilling fluid), 0.5%, 1.0%, and 2.0% by weight according to the
procedures outlined in this section using the Reference Crude Oil:
a. Label 4 jars with the following identification: Jar 1—0%Ref-IOLab,
Jar 2—0.5%Ref-IOLab, Jar 3—l%Ref-IOLab, and Jar 4—2%Ref-
lOLab.
b. Weigh 4, 50-g aliquots of well mixed IO Lab drilling fluid into each of
the 4 jars.
c. Add Reference Oil at 0.5%, 1.0%, and 2.0% by weight to jars 2, 3, and
4 respectively. Jar 1 shall not be spiked with Reference Oil in order to
retain a "0%" oil concentration.
d. Thoroughly mix the contents of each of the 4 jars, using clean glass
stirring rods.
e. Transfer (weigh) a 30-g aliquot from Jar 1 to a labeled centrifuge tube.
Centrifuge the aliquot for a minimum of 15 min at approximately
15,000 rpm, in order to obtain a solids free supernate. Weigh 0.5 g of
the supernate directly into a tared and appropriately labeled GC
straight vial. Spike the 0.5-g supernate with 500 jiL of the O.Olg/mL
1,3,5-trichlorobenzene internal standard solution (see Section 7.2.3),
cap with a Teflon lined crimp cap, and vortex for ca. 10 sec.
f. Repeat step 6.7.2.5(e) except use an aliquot from Jar 2.
g. Repeat step 6.7.2.5(e) except use an aliquot from Jar 3.
6-5
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h. Repeat step 6.7.2.5(e) except use an aliquot from Jar 4.
i. These 4 crude/oil drilling fluid calibration standards are now used for
qualitative and quantitative GC/MS analysis.
6.7.2.6 Precision and recovery standard (mid level crude oil/drilling fluid
calibration standard)—Prepare a midpoint crude oil/ drilling fluid
calibration using IO Lab drilling fluid and Reference Oil at a
concentration of 1.0% by weight. Prepare this standard according to the
procedures outlined in Section 6.7.2.5a through 6.7.2.5e, with the
exception that only "Jar 3" needs to be prepared. Remove and spike with
internal standard, as many 0.5-g aliquots as needed to complete the
GC/MS analysis (see Section 6.11.6—bracketing authentic samples every
12 hours with precision and recovery standard) and the initial
demonstration exercise described in Section 6.9.2.
6.7.2.7 Stability of standards
a. When not used, standards shall be stored in the dark, at -5 to -20°C in
screw-capped vials with PTFE-lined lids. Place a mark on the vial at
the level of the solution so that solvent loss by evaporation can be
detected. Bring the vial to room temperature prior to use.
b. Solutions used for quantitative purposes shall be analyzed within 48
hours of preparation and on a monthly basis thereafter for signs of
degradation. A standard shall remain acceptable if the peak area
remains within ±15% of the area obtained in the initial analysis of the
standard.
6.8 Sample Collection Preservation and Storage
6.8.1 Collect NAF and base fluid samples in 100- to 200-mL glass bottles with PTFE- or
aluminum foil lined caps.
6.8.2 Samples collected in the field shall be stored refrigerated until time of preparation.
6.8.3 Sample and extract holding times for this method have not yet been established.
However, based on initial experience with the method, samples should be analyzed
within seven to ten days of collection and extracts should be analyzed within seven
days of preparation.
6.8.4 After completion of GC/MS analysis, extracts shall be refrigerated at 4°C until further
notification of sample disposal.
6.9 Quality Control
6.9.1 Each laboratory that uses this method is required to operate a formal quality
assurance program (Section 6.16, Reference 4). The minimum requirements of this
program shall consist of an initial demonstration of laboratory capability, and ongoing
analysis of standards, and blanks as a test of continued performance, analyses of
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spiked samples to assess accuracy and analysis of duplicates to assess precision.
Laboratory performance shall be compared to established performance criteria to
determine if the results of analyses meet the performance characteristics of the
method.
6.9.1.1 The analyst shall make an initial demonstration of the ability to generate
acceptable accuracy and precision with this method. This ability shall be
established as described in Section 6.9.2.
6.9.1.2 The analyst is permitted to modify this method to improve separations or
lower the cost of measurements, provided all performance requirements
are met. Each time a modification is made to the method, the analyst is
required to repeat the calibration (Section 6.10.4) and to repeat the initial
demonstration procedure described in Section 6.9.2.
6.9.1.3 Analyses of blanks are required to demonstrate freedom from
contamination. The procedures and criteria for analysis of a blank are
described in Section 6.9.3.
6.9.1.4 Analysis of a matrix spike sample is required to demonstrate method
accuracy. The procedure and QC criteria for spiking are described in
Section 6.9.4.
6.9.1.5 Analysis of a duplicate field sample is required to demonstrate method
precision. The procedure and QC criteria for duplicates are described in
Section 6.9.5.
6.9.1.6 Analysis of a sample of the clean NAF(s) (as sent from the supplier—has
not been circulated downhole) used in the drilling operations is required.
6.9.1.7 The laboratory shall, on an ongoing basis, demonstrate through calibration
verification and the analysis of the precision and recovery standard
(Section 6.7.2.6 ) that the analysis system is in control. These procedures
are described in Section 6.11.6.
6.9.1.8 The laboratory shall maintain records to define the quality of data that is
generated.
6.9.2 Initial precision and accuracy—The initial precision and recovery test shall be
performed using the precision and recovery standard (1% by weight Reference Oil in
IO Lab drilling fluid). The laboratory shall generate acceptable precision and
recovery by performing the following operations.
6.9.2.1 Prepare four separate aliquots of the precision and recovery standard using
the procedure outlined in Section 6.7.2.6 . Analyze these aliquots using the
procedures outlined in Section 6.11.
6-7
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6.9.2.2 Using the results of the set of four analyses, compute the average recovery
(X) in weight percent and the standard deviation of the recovery(s) for
each sample.
6.9.2.3 If s and X meet the acceptance criteria of 80% to 110%, system
performance is acceptable and analysis of samples may begin. If, however,
s exceeds the precision limit or X falls outside the range for accuracy,
system performance is unacceptable. In this event, review this method,
correct the problem, and repeat the test.
6.9.2.4 Accuracy and precision — The average percent recovery (P) and the
standard deviation of the percent recovery (Sp) Express the accuracy
assessment as a percent recovery interval from P-2Spto P+2Sp. For
example, if P=90% and Sp=10% for four analyses of crude oil in NAF, the
accuracy interval is expressed as 70% to 110%. Update the accuracy
assessment on a regular basis.
6.9.3 Blanks — Rinse glassware and centrifuge tubes used in the method with 30 mL of
methylene chloride, remove a 0.5-g aliquot of the solvent, spike it with the 500 jiL of
the internal standard solution (Section 6.7.2.3) and analyze a l-|iL aliquot of the
blank sample using the procedure in Section 6.11. Compute results per Section 6. 12.
6.9.4 Matrix spike sample — Prepare a matrix spike sample according to procedure outlined
in Section 6.7.2.6. Analyze the sample and calculate the concentration (% oil) in the
drilling fluid and % recovery of oil from the spiked drilling fluid using the methods
described in Sections 6.11 and 6. 12.
6.9.5 Duplicates — A duplicate field sample shall be prepared and analyzed according to
Section 6.1 1. The relative percent difference (RPD) of the calculated concentrations
shall be less than 15%.
6.9.5.1 Analyze each of the duplicates per the procedure in Section 6.11 and
compute the results per Section 6. 12.
6.9.5.2 Calculate the relative percent difference (RPD) between the two results
per the following equation:
Where:
DI = Concentration of crude oil in the sample; and
D2 = Concentration of crude oil in the duplicate sample.
6.9.5.3 If the RPD criteria are not met, the analytical system shall be judged to be
out of control, and the problem must be immediately identified and
corrected, and the sample batch re-analyzed.
6-8
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6.9.6 A clean NAF sample shall be prepared and analyzed according to Section 6.11.
Ultimately the oil-equivalent concentration from the TIC or EIP signal measured in
the clean NAF sample shall be subtracted from the corresponding authentic field
samples in order to calculate the true contaminant concentration (% oil) in the field
samples (see Section 6.12).
6.9.7 The specifications contained in this method can be met if the apparatus used is
calibrated properly, and maintained in a calibrated state. The standards used for initial
precision and recovery (Section 6.9.2 ) and ongoing precision and recovery (Section
6.11.6) shall be identical, so that the most precise results will be obtained. The
GC/MS instrument will provide the most reproducible results if dedicated to the
setting and conditions required for the analyses given in this method.
6.9.8 Depending on specific program requirements, field replicates and field spikes of
crude oil into samples may be required when this method is used to assess the
precision and accuracy of the sampling and sample transporting techniques.
6.10 Calibration
6.10.1 Establish gas chromatographic/mass spectrometer operating conditions given in Table
6-1. Perform the GC/MS system hardware-tune as outlined by the manufacture. The
gas chromatograph shall be calibrated using the internal standard technique.
Note: Because each GC is slightly different, it may be necessary to adjust the
operating conditions (carrier gas flow rate and column temperature and temperature
program) slightly until the retention times in Table 6-2 are met.
Table 6-1. Gas Chromatograph/Mass Spectrometer (GC/MS) Operation Conditions
Parameter
Injection port
Transfer line
Detector
Initial Temperature
Initial Time
Ramp
Final Temperature
Final Hold
Carrier Gas
Flow rate
Split ratio
Mass range
Setting
280°C
280°C
280°C
50°C
5 minutes
50 to 300°C @ 5°C per minute
300°C
20 minutes or until all peaks have eluted
Helium
As required for standard operation
As required to meet performance criteria (-1:100)
35 to 600 amu.
6-9
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Table 6-2. Approximate Retention Time for Compounds
Compound
Toluene
Octane, n-C8
Ethylbenzene
1 ,2,4-Trimethylbenzene
Decane, -C10
TCB (Internal Standard)
Dodecane, -C\2
I -Methylnaphthalene
1-Tetradecene
Tetradecane, -Ci4
1,3 -Dimethylnaphthalene
Approximate retention time (minutes)
5.6
7.2
10.3
16.0
16.1
21.3
22.9
26.7
28.4
28.7
29.7
6.10.2 Internal standard calibration procedure—1,3,5-trichlorobenzene (TCB) has been
shown to be free of interferences from diesel and crude oils and is a suitable internal
standard.
6.10.3 The system performance test mix standards prepared in Section 6.7.2.4 shall be used
to establish retention times and establish qualitative detection limits.
6.10.3.1 Spike a 500-mL aliquot of the 1.25 mg/mL SPTM standard with 500 |iL
of the TCB internal standard solution.
6.10.3.2 Inject 1.0 |iL of this spiked SPTM standard onto the GC/MS in order to
demonstrate proper retention times. For the GC/MS used in the
development of this method the ten compounds in the mixture had typical
retention times shown in Table 6-2 above. Extracted ion scans for m/z 91
and 105 showed a maximum abundance of 400,000.
6.10.3.3 Spike a 500-mL aliquot of the 0.125 mg/mL SPTM standard with 500 |iL
of the TCB internal standard solution.
6.10.3.4 Inject 1.0 |iL of this spiked SPTM standard onto the GC/MS to monitor
detectable levels. For the GC/MS used in the development of this test, all
ten compounds showed a minimum peak height of three times signal to
noise. Extracted ion scans for m/z 91 and 105 showed a maximum
abundance of 40,000.
6.10.4 GC/MS crude oil/drilling fluid calibration—There are two methods of quantification:
Total Area Integration (Cg-Co) and EIP Area Integration using m/z's 91 and 105. The
Total Area Integration method should be used as the primary technique for
quantifying crude oil in NAFs. The EIP Area Integration method should be used as a
confirmatory technique for NAFs. However, the EIP Area Integration method shall be
used as the primary method for quantifying oil in enhanced mineral oil (EMO) based
drilling fluid. Inject 1.0 jiL of each of the four crude oil/drilling fluid calibration
6-10
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standards prepared in Section 6.7.2.5 into the GC/MS. The internal standard should
elute approximately 21-22 minutes after injection. For the GC/MS used in the
development of this method, the internal standard peak was (35 to 40)% of full scale
at an abundance of about 3.5e+07.
6.10.4.1 Total Area Integration Method—For each of the four calibration standards
obtain the following: Using a straight baseline integration technique,
obtain the total ion chromatogram (TIC) area from Cg to Co. Obtain the
TIC area of the internal standard (TCB). Subtract the TCB area from the
Cg-Coarea to obtain the true Cg-Co area. Using the Cg-Co and TCB
areas, and known internal standard concentration, generate a linear
regression calibration using the internal standard method. The r2 value for
the linear regression curve shall be greater than or equal to 0.998. Some
synthetic fluids might have peaks that elute in the window and would
interfere with the analysis. In this case the integration window can be
shifted to other areas of scan where there are no interfering peaks from the
synthetic base fluid.
6.10.4.2 EIP Area Integration—For each of the four calibration standards generate
Extracted Ion Profiles (EIPs) for m/z 91 and 105. Using straight baseline
integration techniques, obtain the following EIP areas:
a. For m/z 91 integrate the area under the curve from approximately 9
minutes to 21-22 minutes, just prior to but not including the internal
standard.
b. For m/z 105 integrate the area under the curve from approximately
10.5 minutes to 26.5 minutes.
c. Obtain the internal standard area from the TCB in each of the four
calibration standards, using m/z 180.
d. Using the EIP areas for TCB, m/z 91 and m/z 105, and the known
concentration of internal standard, generate linear regression
calibration curves for the target ions 91 and 105 using the internal
standard method. The r2 value for each of the EIP linear regression
curves shall be greater than or equal to 0.998.
e. Some base fluids might produce a background level that would show
up on the extracted ion profiles, but there should not be any real peaks
(signal to noise ratio of 1:3) from the clean base fluids.
6.11 Procedure
6.11.1 Sample Preparation
6.11.1.1 Mix the authentic field sample (drilling fluid) well. Transfer (weigh) a 30-
g aliquot of the sample to a labeled centrifuge tube.
6-11
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6.11.1.2 Centrifuge the aliquot for a minimum of 15 min at approximately 15,000
rpm, in order to obtain a solids free supernate.
6.11.1.3 Weigh 0.5 g of the supernate directly into a tared and appropriately
labeled GC straight vial.
6.11.1.4 Spike the 0.5-g supernate with 500 |iL of the O.Olg/mL 1,3,5-
trichlorobenzene internal standard solution (see Section 6.7.2.3 ), cap with
a Teflon lined crimp cap, and vortex for ca. 10 sec.
6.11.1.5 The sample is ready for GC/MS analysis.
6.11.2 Gas Chromatography
Table 6-1 summarizes the recommended operating conditions for the GC/MS.
Retention times for the n-alkanes obtained under these conditions are given in Table
6-2. Other columns, chromatographic conditions, or detectors may be used if initial
precision and accuracy requirements (Section 6.9.2 ) are met. The system shall be
calibrated according to the procedures outlined in Section 6.10, and verified every 12
hours according to Section 6.11.6.
6.11.2.1 Samples shall be prepared (extracted) in a batch of no more than 20
samples. The batch shall consist of 20 authentic samples, 1 blank (Section
6.9.3), 1 matrix spike sample (Section 6.9.4), and 1 duplicate field sample
(6.9.5), and a prepared sample of the corresponding clean NAF used in the
drilling process.
6.11.2.2 An analytical sequence shall be analyzed on the GC/MS where the 3
SPTM standards (Section 6.7.2.4) containing internal standard are
analyzed first, followed by analysis of the four GC/MS crude oil/drilling
fluid calibration standards (Section 6.7.2.5), analysis of the blank, matrix
spike sample, the duplicate sample, the clean NAF sample, followed by
the authentic samples.
6.11.2.3 Samples requiring dilution due to excessive signal shall be diluted using
methylene chloride.
6.11.2.4 Inject 1.0 jiL of the test sample or standard into the GC, using the
conditions in Table 6-1.
6.11.2.5 Begin data collection and the temperature program at the time of injection.
6.11.2.6 Obtain a TIC and EIP fingerprint scans of the sample (Table 6-3).
6.11.2.7 If the area of the Cg to Co peaks exceeds the calibration range of the
system, dilute a fresh aliquot of the test sample weighing 0.50-g and re-
analyze.
6-12
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6.11.2.8 Determine the Cg to Co TIC area, the TCB internal standard area, and the
areas for the m/z 91 and 105 EIPs. These shall be used in the calculation
of oil concentration in the samples (see Section 6.12).
Table 6-3. Recommended Ion Mass Numbers
Selected Ion Mass Numbers
91
105
156
Corresponding Aromatic Compounds
Methylbenzene
Ethylbenzene
1 ,4-Dimethylbenzene
1 ,3 -Dimethylbenzene
1 ,2-Dimethylbenzene
1 ,3 ,5 -Trimethylbenzene
1,2,4-Trimethylbenzene
1 ,2,3 -Trimethylbenzene
2,6-Dimethylnaphthalene
1 ,2-Dimethylnaphthalene
1 ,3 -Dimethylnaphthalene
Typical Retention Time (Minutes)
6.0
10.3
10.9
10.9
11.9
15.1
16.0
17.4
28.9
29.4
29.7
6.11.2.9 Observe the presence of peaks in the EIPs that would confirm the presence
of any target aromatic compounds. Using the EIP areas and EIP linear
regression calibrations determine approximate crude oil contamination in
the sample for each of the target ions.
6.11.3 Qualitative Identification—See Section 6.17 of this method for schematic flowchart.
6.11.3.1 Qualitative identification shall be accomplished by comparison of the TIC
and EIP area data from an authentic sample to the TIC and EIP area data
from the calibration standards (see Section 6.10.4). Crude oil shall be
identified by the presence of Cio to Co n-alkanes and corresponding target
aromatics.
6.11.3.2 Using the calibration data, establish the identity of the Cg to Co peaks in
the chromatogram of the sample. Using the calibration data, establish the
identity of any target aromatics present on the extracted ion scans.
6.11.3.3 Crude oil is not present in a detectable amount in the sample if there are
no target aromatics seen on the extracted ion scans. The experience of the
analyst shall weigh heavily in the determination of the presence of peaks
at a signal-to-noise ratio of 3 or greater.
6.11.3.4 If the chromatogram shows n-alkanes from Cg to Co and target aromatics
to be present, contamination by crude oil or diesel shall be suspected and
quantitative analysis shall be determined. If there are no n-alkanes present
that are not seen on the blank, and no target aromatics are seen, the sample
can be considered to be free of contamination.
6-13
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6.11.4 Quantitative Identification
6.11.4.1 Determine the area of the peaks from Cg to Co as outlined in the
calibration section (Section 6.10.4.1). If the area of the peaks for the
sample is greater than that for the clean NAF (base fluid) use the crude
oil/drilling fluid calibration TIC linear regression curve to determine
approximate crude oil contamination.
6.11.4.2 Using the EIPs outlined in Section 6.10.4.2, determine the presence of any
target aromatics. Using the integration techniques outlined in Section
6.10.4.2, obtain the EIP areas for m/z 91 and 105. Use the crude
oil/drilling fluid calibration EIP linear regression curves to determine
approximate crude oil contamination.
6.11.5 Complex Samples
6.11.5.1 The most common interferences in the determination of crude oil can be
from mineral oil, diesel oil, and proprietary additives in drilling fluids.
6.11.5.2 Mineral oil can typically be identified by its lower target aromatic content,
and narrow range of strong peaks.
6.11.5.3 Diesel oil can typically be identified by low amounts of n-alkanes from C?
to Cg, and the absence of n-alkanes greater than C25.
6.11.5.4 Crude oils can usually be distinguished by the presence of high aromatics,
increased intensities of Cg to Co peaks, and/ or the presence of higher
hydrocarbons of Czs and greater (which may be difficult to see in some
synthetic fluids at low contamination levels).
a. Oil condensates from gas wells are low in molecular weight and will
normally produce strong chromatographic peaks in the Cg-Co range.
If a sample of the gas condensate crude oil from the formation is
available, the oil can be distinguished from other potential sources of
contamination by using it to prepare a calibration standard.
b. Asphaltene crude oils with API gravity < 20 may not produce
chromatographic peaks strong enough to show contamination at levels
of the calibration. Extracted ion peaks should be easier to see than
increased intensities for the Cg to Co peaks. If a sample of asphaltene
crude from the formation is available, a calibration standard shall be
prepared.
6.11.6 System and Laboratory Performance
6.11.6.1 At the beginning of each 8-hour shift during which analyses are
performed, GC crude oil/drilling fluid calibration and system performance
test mixes shall be verified. For these tests, analysis of the medium-level
calibration standard (1-% Reference Oil in IO Lab drilling fluid, and 1.25
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mg/mL SPTM with internal standard) shall be used to verify all
performance criteria. Adjustments and/or re-calibration (per Section 6.10)
shall be performed until all performance criteria are met. Only after all
performance criteria are met may samples and blanks be analyzed.
6.11.6.2 Inject 1.0 jiL of the medium-level GC/MS crude oil/drilling fluid
calibration standard into the GC instrument according to the procedures in
Section 6.11.2. Verify that the linear regression curves for both TIC area
and EIP areas are still valid using this continuing calibration standard.
6.11.6.3 After this analysis is complete, inject 1.0 jiL of the 1.25 mg/mL SPTM
(containing internal standard) into the GC instrument and verify the proper
retention times are met (see Table 6-2).
6.11.6.4 Retention times—Retention time of the internal standard. The absolute
retention time of the TCB internal standard shall be within the range 21.0
±0.5 minutes. Relative retention times of the n-alkanes: The retention
times of the n-alkanes relative to the TCB internal standard shall be
similar to those given in Table 6-2.
6.12 Calculations
The concentration of oil in NAFs drilling fluids shall be computed relative to peak areas between
Cg and Co (using the Total Area Integration method) or total peak areas from extracted ion
profiles (using the Extracted Ion Profile Method). In either case, there is a measurable amount of
peak area, even in clean drilling fluid samples, due to spurious peaks and electrometer "noise"
that contributes to the total signal measured using either of the quantification methods. In this
procedure, a correction for this signal is applied, using the blank or clean sample correction
technique described in American Society for Testing Materials (ASTM) Method D-3328-90,
Comparison of Waterborne Oil by Gas Chromatography. In this method, the "oil equivalents"
measured in a blank sample by total area gas chromatography are subtracted from that
determined for a field sample to arrive at the most accurate measure of oil residue in the
authentic sample.
6.12.1 Total Area Integration Method
6.12.1.1 Using Cg to Co TIC area, the TCB area in the clean NAF sample and the
TIC linear regression curve, compute the oil equivalent concentration of
the Cg to Co retention time range in the clean NAF.
Note: The actual TIC area of the Cgto Co is equal to the Cgto Co area
minus the area of the TCB.
6.12.1.2 Using the corresponding information for the authentic sample, compute
the oil equivalent concentration of the Cg to Co retention time range in the
authentic sample.
6-15
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6.12.1.3 Calculate the concentration (% oil) of oil in the sample by subtracting the
oil equivalent concentration (% oil) found in the clean NAF from the oil
equivalent concentration (% oil) found in the authentic sample.
6.12.2 EIP Area Integration Method
6.12.2.1 Using either m/z 91 or 105 EIP areas, the TCB area in the clean NAF
sample, and the appropriate EIP linear regression curve, compute the oil
equivalent concentration of the in the clean NAF.
6.12.2.2 Using the corresponding information for the authentic sample, compute its
oil equivalent concentration.
6.12.2.3 Calculate the concentration (% oil) of oil in the sample by subtracting the
oil equivalent concentration (% oil) found in the clean NAF from the oil
equivalent concentration (% oil) found in the authentic sample.
6.13 Method Performance
6.13.1 Specifications in this method are adopted from EPA Method 1663, Differentiation of
Diesel and Crude Oil by GC/FID (Section 6.16, Reference 4).
6.13.2 Single laboratory method performance using an Internal Olefm (IO) drilling fluid
fortified at 0.5% oil using a 35 API gravity oil was:
Precision and accuracy 94 ±4%
Accuracy interval—86.3% to 102%
Relative percent difference in duplicate analysis—6.2%
6.14 Pollution Prevention
6.14.1 The solvent used in this method poses little threat to the environment when recycled
and managed properly.
6.15 Waste Management
6.15.1 It is the laboratory's responsibility to comply with all federal, state, and local
regulations governing waste management, particularly the hazardous waste
identification rules and land disposal restriction, and to protect the air, water, and land
by minimizing and controlling all releases from fume hoods and bench operations.
Compliance with all sewage discharge permits and regulations is also required.
6.15.2 All authentic samples (drilling fluids) failing the RPE (fluorescence) test (indicated
by the presence of fluorescence) shall be retained and classified as contaminated
samples. Treatment and ultimate fate of these samples is not outlined in this SOP.
6.15.3 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
NW, Washington, DC 20036.
6.16 References
1. Carcinogens—"Working With Carcinogens." Department of Health, Education, and
Welfare, Public Health Service, Centers for Disease Control (available through National
Technical Information Systems, 5285 Port Royal Road, Springfield, VA 22161,
document no. PB-277256): August 1977.
2. "OSHA Safety and Health Standards, General Industry [29 CFR 1910], Revised."
Occupational Safety and Health Administration, OSHA 2206. Washington, DC: January
1976.
3. "Handbook of Analytical Quality Control in Water and Wastewater Laboratories."
USEPA, EMSSL-CI, EPA-600/4-79-019. Cincinnati, OH: March 1979.
4. "Method 1663, Differentiation of Diesel and Crude Oil by GC/FID, Methods for the
Determination of Diesel, Mineral, and Crude Oils in Offshore Oil and Gas Industry
Discharges, EPA 821-R-92-008, Office of Water Engineering and Analysis Division,
Washington, DC: December 1992.
5. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category. Federal Register, 66:
6849(22 January 2001).
6. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category: Correction. Federal
Register, 66: 30811 (8 June 2001).
6-17
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6.17 Schematic Flowchart for Qualitative Identification
Prepare Sample for Analyses
Section 6.11.1
GC/MS Analyses
Obtain the TIC for the Sample
Section 6.11.2
I
No
Determine the Cg to Co TIC Area
for comparison to calibration
standards
Section 6.11.2.8
Obtain the EIP for
m/z 91, m/z 105, and m/z 156
Section 6.11.2.6
I
Crude oil contamination is below
detection limit.
Report not detected
Section 6.11.3.3
Peaks present for
Target Aromatics on
EIP
I
Yes
Integrate peaks on the EIP for comparison
to calibration to determine approximate
crude oil contamination
Section 6.11.4.2
Figure 6-1. Schematic Flowchart for Qualitative Identification
6-18
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7. REVERSE PHASE EXTRACTION (RPE) METHOD FOR DETECTION OF OIL
CONTAMINATION IN NON-AQUEOUS DRILLING FLUIDS (NAF) (EPA METHOD 1670)
7.1 Scope and Application
7.1.1 This method is used for determination of crude or formation oil, or other petroleum
oil contamination, in non-aqueous drilling fluids (NAFs).
7.1.2 This method is intended as a positive/negative test to determine a presence of crude
oil in NAF prior to discharging drill cuttings from offshore production platforms.
7.1.3 This method is for use in the Environmental Protection Agency's (EPA's) survey and
monitoring programs under the Clean Water Act, including monitoring of compliance
with the Gulf of Mexico NPDES General Permit for monitoring of oil contamination
in drilling fluids.
7.1.4 This method has been designed to show positive contamination for 5% of
representative crude oils at a concentration of 0.1% in drilling fluid (vol/vol), 50% of
representative crude oils at a concentration of 0.5%, and 95% of representative crude
oils at a concentration of 1%.
7.1.5 Any modification of this method, beyond those expressly permitted, shall be
considered a major modification subject to application and approval of alternate test
procedures under 40 CFR Parts 136.4 and 136.5.
7.1.6 Each laboratory that uses this method must demonstrate the ability to generate
acceptable results using the procedure in Section 7.9.2.
7.2 Summary of Method
7.2.1 An aliquot of drilling fluid is extracted using isopropyl alcohol.
7.2.2 The mixture is allowed to settle and then filtered to separate out residual solids.
7.2.3 An aliquot of the filtered extract is charged onto a reverse phase extraction (RPE)
cartridge.
7.2.4 The cartridge is eluted with isopropyl alcohol.
7.2.5 Crude oil contaminates are retained on the cartridge and their presence (or absence) is
detected based on observed fluorescence using a black light.
7.3 Definitions
7.3.1 A NAF is one in which the continuous phase is a water immiscible fluid such as an
oleaginous material (e.g., mineral oil, enhance mineral oil, paraffmic oil, or synthetic
material such as olefins and vegetable esters).
7-1
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7.4 Interferences
7.4.1 Solvents, reagents, glassware, and other sample-processing hardware may yield
artifacts that affect results. Specific selection of reagents and purification of solvents
may be required.
7.4.2 All materials used in the analysis shall be demonstrated to be free from interferences
under the conditions of analysis by running laboratory reagent blanks as described in
Section 7.9.5.
7.5 Safety
7.5.1 The toxicity or carcinogenicity of each reagent used in this method has not been
precisely determined; however, each chemical shall be treated as a potential health
hazard. Exposure to these chemicals should be reduced to the lowest possible level.
Material Safety Data Sheets (MSDSs) shall be available for all reagents.
7.5.2 Isopropyl alcohol is flammable and should be used in a well-ventilated area.
7.5.3 Unknown samples may contain high concentration of volatile toxic compounds.
Sample containers should be opened in a hood and handled with gloves to prevent
exposure. In addition, all sample preparation should be conducted in a well-ventilated
area to limit the potential exposure to harmful contaminants. Drilling fluid samples
should be handled with the same precautions used in the drilling fluid handling areas
of the drilling rig.
7.5.4 This method does not address all safety issues associated with its use. The laboratory
is responsible for maintaining a safe work environment and a current awareness file
of OSHA regulations regarding the safe handling of the chemicals specified in this
method. A reference file of material safety data sheets (MSDSs) shall be available to
all personnel involved in these analyses. Additional information on laboratory safety
can be found in Section 7.16, References 1 and 2.
7.6 Equipment and Supplies
Note: Brand names, suppliers, and part numbers are for illustrative purposes only. No
endorsement is implied. Equivalent performance may be achieved using apparatus and materials
other than those specified here, but demonstration of equivalent performance that meets the
requirements of this method is the responsibility of the laboratory.
7.6.1 Sampling equipment.
7.6.1.1 Sample collection bottles/jars—New, pre-cleaned bottles/jars, lot-certified
to be free of artifacts. Glass preferable, plastic acceptable, wide mouth
approximately 1-L, with Teflon-lined screw cap.
7.6.2 Equipment for glassware cleaning.
7.6.2.1 Laboratory sink.
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7.6.2.2 Oven—Capable of maintaining a temperature within ±5°C in the range of
100-250°C.
7.6.3 Equipment for sample extraction.
7.6.3.1 Vials—Glass, 25 mL and 4 mL, with Teflon-lined screw caps, baked at
200-250°C for 1-h minimum prior to use.
7.6.3.2 Gas-tight syringes—Glass, various sizes, 0.5 mL to 2.5 mL (if spiking of
drilling fluids with oils is to occur).
7.6.3.3 Auto pipetters—various sizes, 0.1 mL, 0.5 mL, 1 to 5 mL delivery, and 10
mL delivery, with appropriate size disposable pipette tips, calibrated to
within ±0.5%.
7.6.3.4 Glass stirring rod.
7.6.3.5 Vortex mixer.
7.6.3.6 Disposable syringes—Plastic, 5 mL.
7.6.3.7 Teflon syringe filter, 25-mm, 0.45 |im pore size—Acrodisc®CR Teflon
(or equivalent).
7.6.3.8 Reverse Phase Extraction Cig Cartridge—Waters Sep-Pak®Plus, Cig
Cartridge, 360 mg of sorbent (or equivalent).
7.6.3.9 SPE vacuum manifold—Supelco Brand, 12 unit (or equivalent). Used as
support for cartridge/syringe assembly only. Vacuum apparatus not
required.
7.6.4 Equipment for fluorescence detection.
7.6.4.1 Black light—UV Lamp, Model UVG 11, Mineral Light Lamp, Shortwave
254 nm, or Longwave 365 nm, 15 volts, 60 Hz, 0.16 amps (or equivalent).
7.6.4.2 Black box—cartridge viewing area. A commercially available ultraviolet
viewing cabinet with viewing lamp, or alternatively, a cardboard box or
equivalent, approximately 14 inch x 7.5 inch x 7.5 inch in size and painted
flat black inside. Lamp positioned in fitted and sealed slot in center on top
of box. Sample cartridges sit in a tray, ca. 6 inches from lamp. Cardboard
flaps cut on top panel and side of front panel for sample viewing and
sample cartridge introduction, respectively.
7.6.4.3 Viewing platform for cartridges. Simple support (hand made vial tray—
black in color) for cartridges so that they do not move during the
fluorescence testing.
7-3
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7.7 Reagents and Standards
1.1.1 Isopropyl alcohol—99% purity.
7.7.2 NAF—Appropriate NAF as sent from the supplier (has not been circulated
downhole). Use the clean NAF corresponding to the NAF being used in the current
drilling operation.
7.7.3 Standard crude oil—NIST SRM 1582 petroleum crude oil.
7.8 Sample Collection, Preservation, and Storage
7.8.1 Collect approximately one liter of representative sample (NAF, which has been
circulated downhole) in a glass bottle or jar. Cover with a Teflon lined cap. To allow
for a potential need to re-analyze and/or re-process the sample, it is recommended
that a second sample aliquot be collected.
7.8.2 Label the sample appropriately.
7.8.3 All samples must be refrigerated at 0-4°C from the time of collection until extraction
(40 CFR Part 136, Table II).
7.8.4 All samples must be analyzed within 28 days of the date and time of collection (40
CFR Part 136, Table II).
7.9 Quality Control
7.9.1 Each laboratory that uses this method is required to operate a formal quality
assurance program (Section 7.16, Reference 3). The minimum requirements of this
program consist of an initial demonstration of laboratory capability, and ongoing
analyses of blanks and spiked duplicates to assess accuracy and precision and to
demonstrate continued performance. Each field sample is analyzed in duplicate to
demonstrate representativeness.
7.9.1.1 The analyst shall make an initial demonstration of the ability to generate
acceptable accuracy and precision with this method. This ability is
established as described in Section 7.9.2.
7.9.1.2 Preparation and analysis of a set of spiked duplicate samples to document
accuracy and precision. The procedure for the preparation and analysis of
these samples is described in Section 7.9.4.
7.9.1.3 Analyses of laboratory reagent blanks are required to demonstrate freedom
from contamination. The procedure and criteria for preparation and
analysis of a reagent blank are described in Section 7.9.5.
7.9.1.4 The laboratory shall maintain records to define the quality of the data that
is generated.
7-4
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7.9.1.5 Accompanying QC for the determination of oil in NAF is required per
analytical batch. An analytical batch is a set of samples extracted at the
same time, to a maximum of 10 samples. Each analytical batch of 10 or
fewer samples must be accompanied by a laboratory reagent blank
(Section 7.9.5), corresponding NAF reference blanks (Section 7.9.6), a set
of spiked duplicate samples blank (Section 7.9.4), and duplicate analysis
of each field sample. If greater than 10 samples are to be extracted at one
time, the samples must be separated into analytical batches of 10 or fewer
samples.
7.9.2 Initial demonstration of laboratory capability. To demonstrate the capability to
perform the test, the analyst shall analyze two representative unused drilling fluids
(e.g., internal olefm-based drilling fluid, vegetable ester-based drilling fluid), each
prepared separately containing 0.1%, 1%, and 2% or a representative oil. Each
drilling fluid/concentration combination shall be analyzed 10 times, and successful
demonstration will yield the following average results for the data set:
0.1% oil—Detected in <20% of samples
1% oil—Detected in >75% of samples
2% oil—Detected in >90% of samples
7.9.3 Sample duplicates.
7.9.3.1 The laboratory shall prepare and analyze (Section 7.11.2 and 7.11.4) each
authentic sample in duplicate, from a given sampling site or, if for
compliance monitoring, from a given discharge.
7.9.3.2 The duplicate samples must be compared versus the prepared
corresponding NAF blank.
7.9.3.3 Prepare and analyze the duplicate samples according to procedures
outlined in Section 7.11.
7.9.3.4 The results of the duplicate analyses are acceptable if each of the results
give the same response (fluorescence or no fluorescence). If the results are
different, sample non-homogenicity issues may be a concern. Prepare the
samples again, ensuring a well-mixed sample prior to extraction. Analyze
the samples once again.
7.9.3.5 If different results are obtained for the duplicate a second time, the
analytical system is judged to be out of control and the problem shall be
identified and corrected, and the samples re-analyzed.
7.9.4 Spiked duplicates—Laboratory prepared spiked duplicates are analyzed to
demonstrate acceptable accuracy and precision.
7.9.4.1 Preparation and analysis of a set of spiked duplicate samples with each set
of no more than 10 field samples is required to demonstrate method
accuracy and precision and to monitor matrix interferences (interferences
-------
caused by the sample matrix). A field NAF sample expected to contain
less than 0.5% crude oil (and documented to not fluoresce as part of the
sample batch analysis) shall be spiked with 1% (by volume) of suitable
reference crude oil and analyzed as field samples, as described in Section
7.11. If no low-level drilling fluid is available, then the unused NAF can
be used as the drilling fluid sample.
7.9.5 Laboratory reagent blanks—Laboratory reagent blanks are analyzed to demonstrate
freedom from contamination.
7.9.5.1 A reagent blank is prepared by passing 4 mL of the isopropyl alcohol
through a Teflon syringe filter and collecting the filtrate in a 4-mL glass
vial. A Sep Pak® Cig cartridge is then preconditioned with 3 mL of
isopropyl alcohol. A 0.5-mL aliquot of the filtered isopropyl alcohol is
added to the syringe barrel along with 3.0 mL of isopropyl alcohol. The
solvent is passed through the preconditioned Sep Pak® cartridge. An
additional 2-mL of isopropyl alcohol is eluted through the cartridge. The
cartridge is now considered the "reagent blank" cartridge and is ready for
viewing (analysis). Check the reagent blank cartridge under the black light
for fluorescence. If the isopropyl alcohol and filter are clean, no
fluorescence will be observed.
7.9.5.2 If fluorescence is detected in the reagent blank cartridge, analysis of the
samples is halted until the source of contamination is eliminated and a
prepared reagent blank shows no fluorescence under a black light. All
samples shall be associated with an uncontaminated method blank before
the results may be reported for regulatory compliance purposes.
7.9.6 NAF reference blanks—NAF reference blanks are prepared from the NAFs sent from
the supplier (NAF that has not been circulated downhole) and used as the reference
when viewing the fluorescence of the test samples.
7.9.6.1 A NAF reference blank is prepared identically to the authentic samples.
Place a 0.1 mL aliquot of the "clean" NAF into a 25-mL glass vial. Add
10 mL of isopropyl alcohol to the vial. Cap the vial. Vortex the vial for
approximately 10 sec. Allow the solids to settle for approximately 15
minutes. Using a 5-mL syringe, draw up 4 mL of the extract and filter it
through a PTFE syringe filter, collecting the filtrate in a 4-mL glass vial.
Precondition a Sep Pak® Cig cartridge with 3 mL of isopropyl alcohol.
Add a 0.5-mL aliquot of the filtered extract to the syringe barrel along
with 3.0 mL of isopropyl alcohol. Pass the extract and solvent through the
preconditioned Sep Pak® cartridge. Pass an additional 2-mL of isopropyl
alcohol through the cartridge. The cartridge is now considered the NAF
blank cartridge and is ready for viewing (analysis). This cartridge is used
as the reference cartridge for determining the absence or presence of
fluorescence in all authentic drilling fluid samples that originate from the
same NAF. That is, the specific NAF reference blank cartridge is put
under the black light along with a prepared cartridge of an authentic
sample originating from the same NAF material. The fluorescence or
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absence of fluorescence in the authentic sample cartridge is determined
relative to the NAF reference cartridge.
7.9.6.2 Positive control solution, equivalent to 1% crude oil contaminated mud
extract, is prepared by dissolving 87 mg of standard crude oil into 10.00
mL of methylene chloride. Then mix 40 jiL of this solution into 10.00 mL
of IPA. Transfer 0.5 mL of this solution into a preconditioned Cig
cartridge, followed by 2 ml of IPA.
7.10 Calibration and Standardization
7.10.1 Calibration and standardization methods are not employed for this procedure.
7.11 Procedure
This method is a screening-level test. Precise and accurate results can be obtained only by strict
adherence to all details.
7.11.1 Preparation of the analytical batch.
7.11.1.1 Bring the analytical batch of samples to room temperature.
7.11.1.2 Using a large glass stirring rod, mix the authentic sample thoroughly.
7.11.1.3 Using a large glass stirring rod, mix the clean NAF (sent from the
supplier) thoroughly.
7.11.2 Extraction.
7.11.2.1 Using an automatic positive displacement pipetter and a disposable pipette
tip transfer 0.1-mL of the authentic sample into a 25-mL vial.
7.11.2.2 Using an automatic pipetter and a disposable pipette tip dispense a 10-mL
aliquot of solvent grade isopropyl alcohol (IPA) into the 25 mL vial.
7.11.2.3 Cap the vial and vortex the vial for ca. 10-15 seconds.
7.11.2.4 Let the sample extract stand for approximately 5 minutes, allowing the
solids to separate.
7.11.2.5 Using a 5-mL disposable plastic syringe remove 4 mL of the extract from
the 25-mL vial.
7.11.2.6 Filter 4 mL of extract through a Teflon syringe filter (25-mm diameter,
0.45 |im pore size), collecting the filtrate in a labeled 4-mL vial.
7.11.2.7 Dispose of thePFTE syringe filter.
7.11.2.8 Using a black permanent marker, label a Sep Pak® Cig cartridge with the
sample identification.
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7.11 .2.9 Place the labeled Sep Pak Cig cartridge onto the head of a SPE vacuum
manifold.
7. 1 1 .2. 10 Using a 5-mL disposable plastic syringe, draw up exactly 3-mL (air free)
of isopropyl alcohol.
7. 1 1 .2. 1 1 Attach the syringe tip to the top of the Cig cartridge.
7.11.2.12 Condition the Cig cartridge with the 3-mL of isopropyl alcohol by
depressing the plunger slowly.
Note: Depress the plunger just to the point when no liquid remains in the
syringe barrel. Do not force air through the cartridge. Collect the eluate in
a waste vial.
7. 1 1 .2. 13 Remove the syringe temporarily from the top of the cartridge, then remove
the plunger, and finally reattach the syringe barrel to the top of the Cig
cartridge.
7.11 .2. 14 Using automatic pipetters and disposable pipette tips, transfer 0.5 mL of
the filtered extract into the syringe barrel, followed by a 3.0-mL transfer of
isopropyl alcohol to the syringe barrel.
7.11.2.15 Insert the plunger and slowly depress it to pass only the extract and
solvent through the preconditioned Cig cartridge.
Note: Depress the plunger just to the point when no liquid remains in the
syringe barrel. Do not force air through the cartridge. Collect the eluate in
a waste vial.
7.11.2.16 Remove the syringe temporarily from the top of the cartridge, then remove
the plunger, and finally reattach the syringe barrel to the top of the Cig
cartridge.
7. 1 1 .2. 17 Using an automatic pipetter and disposable pipette tip, transfer 2.0 mL of
isopropyl alcohol to the syringe barrel.
7. 1 1 .2. 1 8 Insert the plunger and slowly depress it to pass the solvent through the Cig
cartridge.
Note: Depress the plunger just to the point when no liquid remains in the
syringe barrel. Do not force air through the cartridge. Collect the eluate in
a waste vial.
7.11.2.19 Remove the syringe and labeled Cig cartridge from the top of the SPE
vacuum manifold.
7.11 .2.20 Prepare a reagent blank according to the procedures outlined in Section
7.9.5.
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7.11.2.21 Prepare the necessary NAF reference blanks for each type of NAF
encountered in the field samples according to the procedures outlined in
Section 7.9.6.
7.11.2.22 Prepare the positive control (1% crude oil equivalent) according to Section
7.9.6.2.
7.11.3 Reagent blank fluorescence testing.
7.11.3.1 Place the reagent blank cartridge in a black box, under a black light.
7.11.3.2 Determine the presence or absence of fluorescence for the reagent blank
cartridge. If fluorescence is detected in the blank, analysis of the samples
is halted until the source of contamination is eliminated and a prepared
reagent blank shows no fluorescence under a black light. All samples must
be associated with an uncontaminated method blank before the results may
be reported for regulatory compliance purposes.
7.11.4 Sample fluorescence testing.
7.11.4.1 Place the respective NAF reference blank (Section 7.9.6) onto the tray
inside the black box.
7.11.4.2 Place the authentic field sample cartridge (derived from the same NAF as
the NAF reference blank) onto the tray, adjacent and to the right of the
NAF reference blank.
7.11.4.3 Turn on the black light.
7.11.4.4 Compare the fluorescence of the sample cartridge with that of the negative
control cartridge (NAF blank, Section 7.9.6.1) and positive control
cartridge (1% crude oil equivalent, Section 7.9.6.2).
7.11.4.5 If the fluorescence of the sample cartridge is equal to or brighter than the
positive control cartridge (1% crude oil equivalent, Section 7.9.6.2 ), the
sample is considered contaminated. Otherwise, the sample is clean.
7.12 Data Analysis and Calculations
Specific data analysis techniques and calculations are not performed in this SOP.
7.13 Method Performance
This method was validated through a single laboratory study, conducted with rigorous statistical
experimental design and interpretation (Section 7.16, Reference 4).
7.14 Pollution Prevention
7.14.1 The solvent used in this method poses little threat to the environment when recycled
and managed properly.
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7.15 Waste Management
7.15.1 It is the laboratory's responsibility to comply with all Federal, State, and local
regulations governing waste management, particularly the hazardous waste
identification rules and land disposal restriction, and to protect the air, water, and land
by minimizing and controlling all releases from bench operations. Compliance with
all sewage discharge permits and regulations is also required.
7.15.2 All authentic samples (drilling fluids) failing the fluorescence test (indicated by the
presence of fluorescence) shall be retained and classified as contaminated samples.
Treatment and ultimate fate of these samples is not outlined in this SOP.
7.15.3 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,
NW, Washington, DC 20036.
7.16 References
1. "Carcinogen—Working with Carcinogens," Department of Health, Education, and
Welfare, Public Health Service, Center for Disease Control, National Institute for
Occupational Safety and Health, Publication No. 77-206, August 1977.
2. "OSHA Safety and Health Standards, General Industry," (29 CFR 1910), Occupational
Safety and Health Administration, OSHA 2206 (Revised, January 1976).
3. "Handbook of Analytical Quality Control in Water and Wastewater Laboratories,"
USEPA, EMSL-Ci, Cincinnati, OH 45268, EPA-600/4-79-019, March 1979.
4. Report of the Laboratory Evaluation of Static Sheen Test Replacements—Reverse Phase
Extraction (RPE) Method for Detecting Oil Contamination in Synthetic Based Mud
(SBM). October 1998. Available from API, 1220 L Street, NW, Washington, DC 20005-
4070, 202-682-8000.
5. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category. Federal Register, 66:
6849(22 January 2001).
6. U.S. EPA. 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category: Correction. Federal
Register, 66: 30811 (8 June 2001).
7-10
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8. DETERMINATION OF THE AMOUNT OF NON-AQUEOUS DRILLING FLUID (NAF) BASE
FLUID FROM DRILL CUTTINGS BY A RETORT CHAMBER (DERIVED FROM API
RECOMMENDED PRACTICE 13B-2) (EPA METHOD 1674)
8.1 Description
8.1.1 This procedure is specifically intended to measure the amount of non-aqueous drilling
fluid (NAF) base fluid from cuttings generated during a drilling operation. This
procedure is a retort test which measures all oily material (NAF base fluid) and water
released from a cuttings sample when heated in a calibrated and properly operating
"Retort" instrument.
8.1.2 In this retort test a known mass of cuttings is heated in the retort chamber to vaporize
the liquids associated with the sample. The NAF base fluid and water vapors are then
condensed, collected, and measured in a precision graduated receiver.
Note: Obtaining a representative sample requires special attention to the details of
sample handling (e.g., location, method, frequency). See Sections 8.5 and 8.6 for
minimum requirements for collecting representative samples. Additional sampling
procedures in a given area may be specified by the NPDES permit controlling
authority.
8.2 Equipment
8.2.1 Retort instrument—The recommended retort instrument has a 50-cm3 volume with an
external heating jacket.
8.2.1.1 Retort Specifications:
a. Retort assembly—retort body, cup and lid.
Material: 303 stainless steel or equivalent.
Volume: Retort cup with lid.
Cup Volume: 50- cm3.
Precision: ±0.25- cm3.
b. Condenser—capable of cooling the oil and water vapors below their
liquification temperature.
c. Heating jacket—nominal 350 watts.
d. Temperature control—capable of limiting temperature of retort to at
least 930°F (500°C) and enough to boil off all NAFs.
8.2.2 Liquid receiver (10- cm3, 20- cm3)—the 10- cm3 and 20- cm3 receivers are specially
designed cylindrical glassware with rounded bottom to facilitate cleaning and funnel-
shaped top to catch falling drops. For compliance monitoring under the NPDES
program, the analyst shall use the 10- cm3 liquid receiver with 0.1 ml graduations to
achieve greater accuracy.
JM
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8.2.2.1 Receiver specifications:
Total volume: 10- cm3, 20- cm3.
Precision (0 to 100%): ±0.05 cm3, ±0.05 cm3.
Outside diameter: 10-mm, 13-mm.
Wall thickness: 1.5±0.1mm, 1.2±0.1mm.
Frequency of graduation marks (0 to 100%): 0.10- cm3, 0.10- cm3.
Calibration: To contain "TC" @ 20°C.
Scale: cm3, cm3.
8.2.2.2 Materi al—Pyrex® or equival ent gl as s.
8.2.3 Toploading balance—capable of weighing 2,000 g and precision of at least 0.1 g.
Unless motion is a problem, the analyst shall use an electronic balance. Where motion
is a problem, the analyst may use a triple beam balance.
8.2.4 Fine steel wool (No. 000)—for packing retort body.
8.2.5 Thread sealant lubricant: high temperature lubricant, e.g. Never-Seez® or equivalent.
8.2.6 Pipe cleaners—to clean condenser and retort stem.
8.2.7 Brush—to clean receivers.
8.2.8 Retort spatula—to clean retort cup.
8.2.9 Corkscrew—to remove spent steel wool.
8.3 Procedure
8.3.1 Clean and dry the retort assembly and condenser.
8.3.2 Pack the retort body with steel wool.
8.3.3 Apply lubricant/sealant to threads of retort cup and retort stem.
8.3.4 Weigh and record the total mass of the retort cup, lid, and retort body with steel wool.
This is mass (A), grams.
8.3.5 Collect a representative cuttings sample (see Note in Section 8.1).
8.3.6 Partially fill the retort cup with cuttings and place the lid on the cup.
8.3.7 Screw the retort cup (with lid) onto the retort body, weigh and record the total mass.
This is mass (B), grams.
8.3.8 Attach the condenser. Place the retort assembly into the heating jacket.
8.3.9 Weigh and record the mass of the clean and dry liquid receiver. This is mass (C),
grams. Place the receiver below condenser outlet.
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8.3.10 Turn on the retort. Allow it to run a minimum of 1 hour.
Note: If solids boil over into receiver, the test shall be rerun. Pack the retort body with
a greater amount of steel wool and repeat the test.
8.3.11 Remove the liquid receiver. Allow it to cool. Record the volume of water recovered.
This is (V), cm3.
Note: If an emulsion interface is present between the oil and water phases, heating the
interface may break the emulsion. As a suggestion, remove the retort assembly from
the heating jacket by grasping the condenser. Carefully heat the receiver along the
emulsion band by gently touching the receiver for short intervals with the hot retort
assembly. Avoid boiling the liquids. After the emulsion interface is broken, allow the
liquid receiver to cool. Read the water volume at the lowest point of the meniscus.
8.3.12 Weigh and record the mass of the receiver and its liquid contents (oil plus water).
This is mass (D), grams.
8.3.13 Turn off the retort. Remove the retort assembly and condenser from the heating jacket
and allow them to cool. Remove the condenser.
8.3.14 Weigh and record the mass of the cooled retort assembly without the condenser. This
is mass (E), grams.
8.3.15 Clean the retort assembly and condenser.
8.4 Calculations
8.4.1 Calculate the mass of oil (NAF base fluid) from the cuttings as follows:
8.4.1.1 Mass of the wet cuttings sample (Mw) equals the mass of the retort
assembly with the wet cuttings sample (B) minus the mass of the empty
retort assembly (A).
MW = B-A [8-1]
8.4.1.2 Mass of the dry retorted cuttings (Mo) equals the mass of the cooled retort
assembly (E) minus the mass of the empty retort assembly (A).
MD = E - A [8-2]
8.4.1.3 Mass of the NAF base fluid (MBp) equals the mass of the liquid receiver
with its contents (D) minus the sum of the mass of the dry receiver (C) and
the mass of the water (V).
MBF = D-(C + V) [8-3]
Note: Assuming the density of water is 1 g/cm3, the volume of water is
equivalent to the mass of the water.
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8.4.2 Mass balance requirement:
The sum of MD, MBF, and V shall be within 5% of the mass of the wet sample.
MD + MBF + V
M
w
= 0.95 to 1.05 [8-4]
The procedure shall be repeated if this requirement is not met.
8.4.3 Reporting oil from cuttings:
8.4.3. 1 Assume that all oil recovered is NAF base fluid.
8.4.3.2 The mass percent NAF base fluid retained on the cuttings (%BF;) for the
sampled discharge "i" is equal to 100 times the mass of the NAF base
fluid (MBF) divided by the mass of the wet cuttings sample (Mw).
%BFi= x 100 [8-5]
1V1W
Operators discharging small volume NAF-cuttings discharges which do
not occur during a NAF-cuttings discharge sampling interval (i.e.,
displaced interfaces, accumulated solids in sand traps, pit clean-out solids,
or centrifuge discharges while cutting mud weight) shall either: (a)
Measure the mass percent NAF base fluid retained on the cuttings
(%BFSvo) for each small volume NAF-cuttings discharges; or (b) use a
default value of 25% NAF base fluid retained on the cuttings.
8.4.3.3 The mass percent NAF base fluid retained on the cuttings is determined
for all cuttings wastestreams and includes fines discharges and any
accumulated solids discharged. (See Section 8.4.3.6 for procedures on
measuring or estimating the mass percent NAF base fluid retained on the
cuttings (%BF) for dual gradient drilling seafloor discharges performed to
ensure proper operation of subsea pumps.)
8.4.3.4 A mass NAF-cuttings discharge fraction (X, unitless) is calculated for all
NAF-cuttings, fines, or accumulated solids discharges every time a set of
retorts is performed (Section 8.4.3.6 for procedures on measuring or
estimating the mass NAF-cuttings discharge fraction (X) for dual gradient
drilling seafloor discharges performed to ensure proper operation of
subsea pumps). The mass NAF-cuttings discharge fraction (X) combines
the mass of NAF-cuttings, fines, or accumulated solids discharged from a
particular discharge over a set period of time with the total mass of NAF-
cuttings, fines, or accumulated solids discharged into the ocean during the
same period of time (see Sections 8.5 and 8.6). The mass NAF-cuttings
discharge fraction (X) for each discharge is calculated by direct
measurement as:
8-4
-------
Xi = ~- [8-6]
Where:
X; = Mass NAF-cuttings discharge fraction for NAF-cuttings, fines, or
accumulated solids discharge "i", (unitless)
F; = Mass of NAF-cuttings discharged from NAF-cuttings, fines, or
accumulated solids discharge "i" over a specified period of time
(see Sections 8.5 and 8.6), (kg)
G = Mass of all NAF-cuttings discharges into the ocean during the
same period of time as used to calculate Fi, (kg)
If an operator has more than one point of NAF-cuttings discharge, the
mass faction (X;) must be determined by: (a) Direct measurement (see
Equation 8-6); (b) using the following default values of 0.85 and 0.15 for
the cuttings dryer (e.g., horizontal centrifuge, vertical centrifuge, squeeze
press, High-G linear shakers) and fines removal unit (e.g., decanting
centrifuges, mud cleaners), respectively, when the operator is only
discharging from the cuttings dryer and the fines removal unit; or (c) using
direct measurement of "F;" (see Equation 8-6) for fines and accumulated
solids, using Equation 8-6A to calculate "GEST" for use as "G" in Equation
8-6, and calculating the mass (kg) of NAF-cuttings discharged from the
cuttings dryer (F;) as the difference between the mass of "GEST" calculated
in Equation 8-6A (kg) and the sum of all fines and accumulated solids
mass directly measured (kg) (see Equation 8-6).
GEST = Estimated mass of all NAF-cuttings discharges into the ocean
during the same period of time as used to calculate F;
(see Equation 8-6), (kg) [8-6A]
Where:
GEST = Hole Volume (bbl) x (396.9 kg/bbl) x (1 + Z/100)
Z = The base fluid retained on cuttings limitation or standard (%)
which apply to the NAF being discharge (see 40 CFR§§435.13.
and 435.15).
Hole Volume = [Cross-Section Area of NAF interval (in2)] x Average
(bbl) Rate of Penetration (feet/hr) x period of time (min) used
to calculate F; (see Equation 8-6) x (1 hr/60 min) x
(Ibbl/5.61ft3)x(lft/12in)2
Cross-Section = (3.14 x [Bit Diameter (in)]2)/4
Area of NAF
r\
interval (in )
Bit Diameter = Diameter of drilling bit for the NAF interval producing
(in) drilling cuttings during the same period of time as used to
calculate F; (see Equation 8-6)
Average Rate = Arithmetic average of rate of penetration into the
of Penetration formation during the same period of time as used to
(feet/hr) calculate F; (see Equation 8-6)
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Note: Operators with one NAF-cuttings discharge may set the mass NAF-
cuttings discharge fraction (Xj) equal to 1.0.
8.4.3.5 Each NAF-cuttings, fines, or accumulated solids discharge has an
associated mass percent NAF base fluid retained on cuttings value (%BF)
and mass NAF-cuttings discharge fraction (X) each time a set of retorts is
performed. A single total mass percent NAF base fluid retained on
cuttings value (%BFi) is calculated every time a set of retorts is
performed. The single total mass percent NAF base fluid retained on
cuttings value (%BFx) is calculated as:
%BFTj=Z(Xi)x(%BFi) [8-7]
Where:
%BFxj = Total mass percent NAF base fluid retained on cuttings value for
retort set "j" (unitless as percentage, %)
X; = Mass NAF-cuttings discharge fraction for NAF-cuttings, fines, or
accumulated solids discharge "i", (unitless)
%BF; = Mass percent NAF base fluid retained on the cuttings for NAF-
cuttings, fines, or accumulated solids discharge "i", (unitless as
percentage, %)
Note:SXj= 1.
Operators with one NAF-cuttings discharge may set %BFx,j equal to
%BFi.
8.4.3.6 Operators performing dual gradient drilling operations may require
seafloor discharges of large cuttings (>l/4') to ensure the proper operation
of subsea pumps (e.g., electrical submersible pumps). Operators
performing dual gradient drilling operations which lead to seafloor
discharges of large cuttings for the proper operation of subsea pumps shall
either: (a) Measure the mass percent NAF base fluid retained on cuttings
value (%BF) and mass NAF-cuttings discharge fraction (X) for seafloor
discharges each time a set of retorts is performed; (b) use the following set
of default values, (%BF = 14%; X = 0.15); or (c) use a combination of (a)
and (b) (e.g., use a default value for %BF and measure X).
Additionally, operators performing dual gradient drilling operations which
lead to seafloor discharges of large cuttings for the proper operation of
subsea pumps shall also perform the following tasks:
a. Use side scan sonar or shallow seismic to determine the presence of
high density chemosynthetic communities. Chemosynthetic
communities are assemblages of tube worms, clams, mussels, and
bacterial mats that occur at natural hydrocarbon seeps or vents,
generally in water depths of 500 meters or deeper. Seafloor discharges
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of large cuttings for the proper operation of subsea pumps shall not be
permitted within 1,000 feet of a high density chemosynthetic
community.
b. Seafloor discharges of large cuttings for the proper operation of subsea
pumps shall be visually monitored and documented by a Remotely
Operated Vehicle (ROV) within the tether limit (approximately 300
feet). The visual monitoring shall be conducted prior to each time the
discharge point is relocated (cuttings discharge hose) and conducted
along the same direction as the discharge hose position. Near-seabed
currents shall be obtained at the time of the visual monitoring.
c. Seafloor discharges of large cuttings for the proper operation of subsea
pumps shall be directed within a 150 foot radius of the wellbore.
8.4.3.7 The weighted mass ratio averaged over all NAF well sections (%BFweu) is
the compliance value that is compared with the "maximum weighted mass
ratio averaged over all NAF well sections" BAT discharge limitations (see
the table in 40 CFR §435.13 and footnote 5 of the table in 40 CFR
§435.43) or the "maximum weighted mass ratio averaged over all NAF
well sections" NSPS discharge limitations (see the table in 40 CFR
§435.15 and footnote 5 of the table in 40 CFR §435.45). The weighted
mass ratio averaged over all NAF well sections (%BFwen) is calculated as
the arithmetic average of all total mass percent NAF base fluid retained on
cuttings values (%BFT) and is given by the following expression:
%BFwell=-
[8.g]
Where:
%BFweii = Weighted mass ratio averaged over all NAF well sections (unitless
as percentage, %)
%BFTj = Total mass percent NAF base fluid retained on cuttings value for
retort set "j" (unitless as percentage, %)
n = Total number of retort sets performed over all NAF well sections
(unitless)
Small volume NAF-cuttings discharges which do not occur during a NAF-
cuttings discharge sampling interval (i.e., displaced interfaces,
accumulated solids in sand traps, pit clean-out solids, or centrifuge
discharges while cutting mud weight) shall be mass averaged with the
arithmetic average of all total mass percent NAF base fluid retained on
cuttings values (see Equation 8-8). An additional sampling interval shall
be added to the calculation of the weighted mass ratio averaged over all
NAF well sections (%BFweii). The mass fraction of the small volume
NAF-cuttings discharges (Xsvo) will be determined by dividing the mass
3-7
-------
Where:
XSVD
FSVD
Gwell
Where:
FSVD
Psvd
VSVD
of the small volume NAF-cuttings discharges (Fsvo) by the total mass of
NAF-cuttings discharges for the well drilling operation (Gweii+ FSVD)-
XSVD -
FSVD
weii
[8-9]
= Mass fraction of the small volume NAF-cuttings discharges
(unitless)
= Mass of the small volume NAF-cuttings discharges (kg)
= Mass of total NAF-cuttings from the well (kg)
The mass of small volume NAF-cuttings discharges (Fsvo) shall be
determined by multiplying the density of the small volume NAF-cuttings
discharges (psvd) times the volume of the small volume NAF-cuttings
discharges (VSvo).
FSVD — Psvd x VSVD
[8-10]
= Mass of small volume NAF-cuttings discharges (kg)
= Density of the small volume NAF-cuttings discharges (kg/bbl)
= Volume of the small volume NAF-cuttings discharges (bbl)
The density of the small volume NAF-cuttings discharges shall be
measured. The volume of small volume discharges (VSVD) shall be either:
(a) Be measured or (b) use default values of 10 bbl of SBF for each
interface loss and 75 bbl of SBM for pit cleanout per well.
The total mass of NAF-cuttings discharges for the well (Gweii) shall be
either: (a) Measured; or (b) calculated by multiplying 1.0 plus the
arithmetic average of all total mass percent NAF base fluid retained on
cuttings values [see Equation 8-8] times the total hole volume (Vweii) for
all NAF well sections times a default value for the density the formation
of 2.5 g/cm3 (396.9 kg/bbl).
1 +
xVwell(bbl)x396.9-^-
bbl
[8-11]
Where:
Gwell
S(%BFT.j)
V
well
= Total mass of NAF-cuttings discharges for the well (kg)
see Equation 8-8 (unitless as a percentage)
Total hole volume (Vweii) for all NAF well sections (bbl)
-------
VWeii (barrels) =
The total hole volume of NAF well sections (Vweii) will be calculated as:
Bit diameter (in)2
-
1 029
. . , , , ,„.
chan§e m measured depth (ft)
,-„,„-,
[8-12]
For wells where small volume discharges associated with cuttings are
made, %BFwen becomes:
(l-XSVD)>
+ XSVD x%BFSVD
[8-13]
8.4.3.8
8.4.3.9
Note: See Sections 8.5 and 8.6 to determine the sampling frequency to
determine the total number of retort sets required for all NAF well
sections.
The total number of retort sets (n) is increased by 1 for each sampling
interval (see Section 8.5.2.4) when all NAF cuttings, fines, or accumulated
solids for that sampling interval are retained for no discharge. A zero
discharge interval shall be at least 500 feet up to a maximum of three per
day. This action has the effect of setting the total mass percent NAF base
fluid retained on cuttings value (%BFT) at zero for that NAF sampling
interval when all NAF cuttings, fines, or accumulated solids are retained
for no discharge.
Operators that elect to use the Best Management Practices (BMPs) for
NAF-cuttings shall use the procedures outlined in Section 8.6.
8.5 Requirements for Sampling Cuttings Discharge Streams for use with this Method
8.5.1 Sampling Locations
8.5.1.1 Each NAF-cuttings waste stream that discharges into the ocean shall be
sampled and analyzed as detailed earlier in Section 8.3. NAF-cuttings
discharges to the ocean may include discharges from primary shakers,
secondary shakers, cuttings dryer, fines removal unit, accumulated solids,
and any other cuttings separation device whose NAF-cuttings waste is
discharged to the ocean. NAF-cuttings wastestreams not directly
discharged to the ocean (e.g., NAF-cuttings generated from shake shakers
and sent to a cuttings dryer for additional processing) do not require
sampling and analysis.
8.5.1.2 The collected samples shall be representative of each NAF-cuttings
discharge. Operators shall conduct sampling to avoid the serious
consequences of error (i.e., bias or inaccuracy). Operators shall collect
NAF-cuttings samples near the point of origin and before the solids and
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liquid fractions of the stream have a chance to separate from one another.
For example, operators shall collect shale shaker NAF-cuttings samples at
the point where NAF-cuttings are coming off the shale shaker and not
from a holding container downstream where separation of larger particles
from the liquid can take place.
8.5.1.3 Operators shall provide a simple schematic diagram of the solids control
system and sample locations to the NPDES permit controlling authority.
8.5.2 Type of Sample and Sampling Frequency
8.5.2.1 Each NAF-cuttings, fines, or accumulated solids discharge has an
associated mass percent NAF base fluid retained on cuttings value (%BF)
and mass NAF-cuttings discharge fraction (X) for each sampling interval
(see Section 8.5.2.4). Operators shall collect a single discrete NAF-
cuttings sample for each NAF-cuttings waste stream discharged to the
ocean during every sampling interval.
8.5.2.2 Operators shall use measured depth in feet from the Kelly bushing when
samples are collected.
8.5.2.3 The NAF-cuttings samples collected for the mass fraction analysis (see
Equation 8-6) shall also be used for the retort analysis (see Equations 8-1
through 8-5).
8.5.2.4 Operators shall collect and analyze at least one set of NAF-cuttings
samples per day while discharging. Operators engaged in fast drilling (i.e.,
greater than 500 linear NAF feet advancement of drill bit per day) shall
collect and analyze one set of NAF-cuttings samples per 500 linear NAF
feet of footage drilled. Operators are not required to collect and analyze
more than three sets of NAF-cuttings samples per day (i.e., three sampling
intervals). Operators performing zero discharge of all NAF-cuttings (i.e.,
all NAF cuttings, fines, or accumulated solids retained for no discharge)
shall use the following periods to count sampling intervals: (1) One
sampling interval per day when drilling is less than 500 linear NAF feet
advancement of drill bit per day; and (2) one sampling interval per 500
linear NAF feet of footage drilled with a maximum of three sampling
intervals per day.
8.5.2.5 The operator shall measure the individual masses (F;, kg) and sum total
mass (G, kg) (see Equation 8-6) over a representative period of time (e.g.,
<10 minutes) during steady-state conditions for each sampling interval
(see Section 8.5.2.4). The operator shall ensure that all NAF-cuttings are
capture for mass analysis during the same sampling time period (e.g., <10
minutes) at approximately the same time (i.e., all individual mass samples
collected within one hour of each other).
8.5.2.6 Operators using Best Management Practices (BMPs) to control NAF-
cuttings discharges shall follow the procedures in Section 8.6.
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8.5.3 Sample Size and Handling
8.5.3.1 The volume of each sample depends on the volumetric flow rate (cm3/s) of
the NAF-cuttings stream and the sampling time period (e.g., <10 minutes).
Consequently, different solids control equipment units producing different
NAF-cuttings waste streams at different volumetric flow rates will
produce different size samples for the same period of time. Operators shall
use appropriately sized sample containers for each NAF-cuttings waste
stream to ensure no NAF-cuttings are spilled during sample collection.
Operators shall use the same time period (e.g., <10 minutes) to collect
NAF-cuttings samples from each NAF-cuttings waste stream. Each NAF-
cuttings sample size shall be at least one gallon. Operators shall clearly
mark each container to identify each NAF-cuttings sample.
8.5.3.2 Operators shall not decant, heat, wash, or towel the NAF-cuttings to
remove NAF base fluid before mass and retort analysis.
8.5.3.3 Operators shall first calculate the mass of each NAF-cuttings sample and
perform the mass ratio analysis (see Equation 8-6). Operators with only
one NAF-cuttings discharge may skip this step (see Section 8.4.3.4).
8.5.3.4 Operators shall homogenize (e.g., stirring, shaking) each NAF-cuttings
sample prior to placing a sub-sample into the retort cup. The bottom of the
NAF-cuttings sample container shall be examined to be sure that solids are
not sticking to it.
8.5.3.5 Operators shall then calculate the NAF base fluid retained on cuttings
using the retort procedure (See Equations 8-1 through 8-5). Operators
shall start the retort analyses no more than two hours after collecting the
first individual mass sample for the sampling interval.
8.5.3.6 Operators shall not discharge any sample before successfully completing
the mass and retort analyses [i.e., mass balance requirements (see Section
8.4.2) are satisfied]. Operators shall immediately re-run the retort analyses
if the mass balance requirements (see Equation 8-4) are not within a
tolerance of 5% (see Section 8.4.2, Equation 8-4).
8.5.4 Calculations
8.5.4.1 Operators shall calculate a set of mass percent NAF base fluid retained on
cuttings values (%BF) and mass NAF-cuttings discharge fractions (X) for
each NAF-cuttings waste stream (see Section 8.5.1.1) for each sampling
interval (see Section 8.5.2.4) using the procedures outlined earlier in
Section 8.4.
8.5.4.2 Operators shall tabulate the following data for each individual NAF-
cuttings sample: (1) Date and time of NAF-cuttings sample collection; (2)
time period of NAF-cuttings sample collection (see Section 8.5.3.1); (3)
mass and volume of each NAF-cuttings sample; (4) measured depth (feet)
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at NAF-cuttings sample collection (see Section 8.5.2.2); (5) respective
linear feet of hole drilled represented by the NAF-cuttings sample (feet);
and (6) the drill bit diameter (inches) used to generate the NAF-cuttings
sample cuttings.
8.5.4.3 Operators shall calculate a single total mass percent NAF base fluid
retained on cuttings value (%BFT) for each sampling interval (see Section
8.5.2.4) using the procedures outlined in Section 8.4.
8.5.4.4 Operators shall tabulate the following data for each total mass percent
NAF base fluid retained on cuttings value (%BFT) for each NAF-cuttings
sampling interval: (1) Date and starting and stopping times of NAF-
cuttings sample collection and retort analyses; (2) measured depth of well
(feet) at start of NAF-cuttings sample collection (see Section 8.5.2.2); (3)
respective linear feet of hole drilled represented by the NAF-cuttings
sample (feet); (4) the drill bit diameter (inches) used to generate the NAF-
cuttings sample cuttings; and (5) annotation when zero discharge of NAF-
cuttings is performed.
8.5.4.5 Operators shall calculate the weighted mass ratio averaged over all NAF
well sections (%BFwen) using the procedures outlined in Section 8.4.
8.5.4.6 Operators shall tabulate the following data for each weighted mass ratio
averaged over all NAF well sections (%BFwen) for each NAF well: (1)
Starting and stopping dates of NAF well sections; (2) measured depth
(feet) of all NAF well sections; (3) total number of sampling intervals (see
Sections 8.5.2.4 and 8.5.2.6); (4) number of sampling intervals tabulated
during any zero discharge operations; (5) total volume of zero discharged
NAF-cuttings over entire NAF well sections; and (6) identification of
whether BMPs were employed (see Section 8.6).
8.6 Best Management Practices (BMPs) for use with this Method
8.6.1 Overview of BMPs
8.6.1.1 Best Management Practices (BMPs) are inherently pollution prevention
practices. BMPs may include the universe of pollution prevention
encompassing production modifications, operational changes, material
substitution, materials and water conservation, and other such measures.
BMPs include methods to prevent toxic and hazardous pollutants from
reaching receiving waters. Because BMPs are most effective when
organized into a comprehensive facility BMP Plan, operators shall develop
a BMP in accordance with the requirements in this addendum.
8.6.1.2 The BMP requirements contained in this section were compiled from
several Regional permits, an EPA guidance document (i.e., Guidance
Document for Developing Best Management Practices (BMP)" (EPA
833-B-93-004, U.S. EPA, 1993)), and draft industry BMPs. These
common elements represent the appropriate mix of broad directions
-------
needed to complete a BMP Plan along with specific tasks common to all
drilling operations.
8.6.1.3 Operators are not required to use BMPs if all NAF-cuttings discharges are
monitored in accordance with Sections 8.1 through 8.4.
8.6.2 BMP Plan Purpose and Objectives
8.6.2.1 Operators shall design the BMP Plan to prevent or minimize the
generation and the potential for the discharge of NAF from the facility to
the waters of the United States through normal operations and ancillary
activities. The operator shall establish specific objectives for the control of
NAF by conducting the following evaluations.
8.6.2.2 The operator shall identify and document each NAF well that uses BMPs
before starting drilling operations and the anticipated total feet to be
drilled with NAF for that particular well.
8.6.2.3 Each facility component or system controlled through use of BMPs shall
be examined for its NAF-waste minimization opportunities and its
potential for causing a discharge of NAF to waters of the United States
due to equipment failure, improper operation, natural phenomena (e.g.,
rain, snowfall).
8.6.2.4 For each NAF wastestream controlled through BMPs where experience
indicates a reasonable potential for equipment failure (e.g., a tank
overflow or leakage), natural condition (e.g., precipitation), or other
circumstances to result in NAF reaching surface waters, the BMP Plan
shall include a prediction of the total quantity of NAF which could be
discharged from the facility as a result of each condition or circumstance.
8.6.3 BMP Plan Requirements
8.6.3.1 The BMP Plan may reflect requirements within the pollution prevention
requirements required by the Minerals Management Service (see 30 CFR
250.300) or other Federal or State requirements and incorporate any part
of such plans into the BMP Plan by reference.
8.6.3.2 The operator shall certify that its BMP Plan is complete, on-site, and
available upon request to EPA or the NPDES Permit controlling authority.
This certification shall identify the NPDES permit number and be signed
by an authorized representative of the operator. This certification shall be
kept with the BMP Plan. For new or modified NPDES permits, the
certification shall be made no later than the effective date of the new or
modified permit. For existing NPDES permits, the certification shall be
made within one year of permit issuance.
8.6.3.3 The BMP Plan shall:
8-13
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a. Be documented in narrative form, and shall include any necessary plot
plans, drawings or maps, and shall be developed in accordance with
good engineering practices. At a minimum, the BMP Plan shall
contain the planning, development and implementation, and
evaluation/reevaluation components. Examples of these components
are contained in "Guidance Document for Developing Best
Management Practices (BMP)" (EPA 833-B-93-004, U.S. EPA,
1993).
b. Include the following provisions concerning BMP Plan review.
• Be reviewed by permittee's drilling engineer and offshore
installation manager (OEVI) to ensure compliance with the BMP
Plan purpose and objectives set forth in Section 8.6.2.
• Include a statement that the review has been completed and that
the BMP Plan fulfills the BMP Plan purpose and objectives set
forth in Section 8.6.2. This statement shall have dated signatures
from the permittee's drilling engineer and offshore installation
manager and any other individuals responsible for development
and implementation of the BMP Plan.
8.6.3.4 Address each component or system capable of generating or causing a
release of significant amounts of NAF and identify specific preventative or
remedial measures to be implemented.
8.6.4 BMP Plan Documentation
8.6.4.1 The operator shall maintain a copy of the BMP Plan and related
documentation (e.g., training certifications, summary of the monitoring
results, records of NAF-equipment spills, repairs, and maintenance) at the
facility and shall make the BMP Plan and related documentation available
to EPA or the NPDES Permit controlling authority upon request.
8.6.5 BMP Plan Modification
8.6.5.1 For those NAF wastestreams controlled through BMPs, the operator shall
amend the BMP Plan whenever there is a change in the facility or in the
operation of the facility which materially increases the generation of those
NAF-wastes or their release or potential release to the receiving waters.
8.6.5.2 At a minimum the BMP Plan shall be reviewed once every five years and
amended within three months if warranted. Any such changes to the BMP
Plan shall be consistent with the objectives and specific requirements
listed in this addendum. All changes in the BMP Plan shall be reviewed by
the permittee's drilling engineer and offshore installation manager.
8.6.5.3 At any time, if the BMP Plan proves to be ineffective in achieving the
general objective of preventing and minimizing the generation of NAF-
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wastes and their release and potential release to the receiving waters
and/or the specific requirements in this addendum, the permit and/or the
BMP Plan shall be subject to modification to incorporate revised BMP
requirements.
8.6.6 Specific Pollution Prevention Requirements for NAF Discharges Associated with
Cuttings
8.6.6.1 The following specific pollution prevention activities are required in a
BMP Plan when operators elect to control NAF discharges associated with
cuttings by a set of BMPs.
8.6.6.2 Establishing programs for identifying, documenting, and repairing
malfunctioning NAF equipment, tracking NAF equipment repairs, and
training personnel to report and evaluate malfunctioning NAF equipment.
8.6.6.3 Establishing operating and maintenance procedures for each component in
the solids control system in a manner consistent with the manufacturer's
design criteria.
8.6.6.4 Using the most applicable spacers, flushes, pills, and displacement
techniques in order to minimize contamination of drilling fluids when
changing from water-based drilling fluids to NAF and vice versa.
8.6.6.5 A daily retort analysis shall be performed (in accordance with Sections 8.1
through 8.4) during the first 0.33 X feet drilled with NAF where X is the
anticipated total feet to be drilled with NAF for that particular well. The
retort analyses shall be documented in the well retort log. The operators
shall use the calculation procedures detailed in Section 8.4 (see Equations
8-1 through 8-8) to determine the arithmetic average (%BFwen) of the
retort analyses taken during the first 0.33 X feet drilled with NAF.
a. When the arithmetic average (%BFwell) of the retort analyses taken
during the first 0.33 X feet drilled with NAF is less than or equal to the
base fluid retained on cuttings limitation or standard (see 40 CFR
§§435.13 and 435.15), retort monitoring of cuttings may cease for that
particular well. The same BMPs and drilling fluid used during the first
0.33 X feet shall be used for all remaining NAF sections for that
particular well.
b. When the arithmetic average (%BFweii) of the retort analyses taken
during the first 0.33 X feet drilled with NAF is greater the base fluid
retained on cuttings limitation or standard (see 40 CFR §§435.13 and
435.15), retort monitoring shall continue for the following (second)
0.33 X feet drilled with NAF where X is the anticipated total feet to be
drilled with NAF for that particular well. The retort analyses for the
first and second 0.33 X feet shall be documented in the well retort log.
8-15
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• When the arithmetic average (%BFweii) of the retort analyses taken
during the first 0.66 X feet (i.e., retort analyses taken from first and
second 0.33 X feet) drilled with NAF is less than or equal to the
base fluid retained on cuttings limitation or standard (see 40 CFR
§§435.13 and 435.15), retort monitoring of cuttings may cease for
that particular well. The same BMPs and drilling fluid used during
the first 0.66 X feet shall be used for all remaining NAF sections
for that particular well.
• When the arithmetic average (%BFweii) of the retort analyses taken
during the first 0.66 X feet (i.e., retort analyses taken from first and
second 0.33 X feet) drilled with NAF is greater than the base fluid
retained on cuttings limitation or standard (see 40 CFR §§435.13
and 435.15), retort monitoring shall continue for all remaining
NAF sections for that particular well. The retort analyses for all
NAF sections shall be documented in the well retort log.
c. When the arithmetic average (%BFwell) of the retort analyses taken
over all NAF sections for the entire well is greater that the base fluid
retained on cuttings limitation or standard (see 40 CFR §§435.13 and
435.15), the operator is in violation of the base fluid retained on
cuttings limitation or standard and shall submit notification of these
monitoring values in accordance with NPDES permit requirements.
Additionally, the operator shall, as part of the BMP Plan, initiate a
reevaluation and modification to the BMP Plan in conjunction with
equipment vendors and/or industry specialists.
d. The operator shall include retort monitoring data and dates of retort-
monitored and non-retort-monitored NAF-cuttings discharges
managed by BMPs in their NPDES permit reports.
8.6.6.6 Establishing mud pit and equipment cleaning methods in such a way as to
minimize the potential for building-up drill cuttings (including
accumulated solids) in the active mud system and solids control equipment
system. These cleaning methods shall include but are not limited to the
following procedures.
a. Ensuring proper operation and efficiency of mud pit agitation
equipment.
b. Using mud gun lines during mixing operations to provide agitation in
dead spaces.
c. Pumping drilling fluids off of drill cuttings (including accumulated
solids) for use, recycle, or disposal before using wash water to
dislodge solids.
8-16
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9. PAH CONTENT OF OIL BY HPLC/UV (EPA METHOD 1654, REVISION A)
9.1 Scope and Application
9.1.1 This method is designed to determine the polynuclear aromatic hydrocarbon (PAH)
content of oil by high-performance liquid chromatography (HPLC) with an ultra-
violet absorption (UV) detector. The PAH content is measured and reported as
phenanthrene.
9.1.2 This method is for use in the Environmental Protection Agency's (EPA's) survey and
monitoring programs under the Federal Water Pollution Control Act.
9.1.3 For oil in drilling muds, this method is designed to be used in conjunction with the
extraction procedure in EPA Method 1662.
9.1.4 The level of PAH in Table 9-1 typifies the minimum level that can be detected in oil
with this method.
9.1.5 Any modification of this method beyond those expressly permitted shall be
considered as a major modification subject to application and approval of alternative
test procedures under 40 CFR 136.4 and 136.5.
9.1.6 This method is restricted to use by or under the supervision of analysts experienced in
the use of HPLC systems and in the interpretation of liquid chromatograms. Each
analyst must demonstrate the ability to generate acceptable results with this method
using the procedure described in Section 9.8.2.
9.2 Summary of Method
9.2.1 An oil sample is diluted in acetonitrile and a 20-uL aliquot is injected into the HPLC.
The PAHs are partially separated by HPLC and detected with the UV detector.
9.2.2 Identification of PAH (qualitative analysis) is performed by comparing the response
of the UV detector to the response during the retention-time range characteristic of
the PAH in diesel oil. PAH is present when a response occurs during this retention-
time range.
9.2.3 Quantitative analysis is performed by calibrating the HPLC with phenanthrene using
an external standard technique, and using the calibration factor to determine the
concentration of PAH in the sample.
9.2.4 Quality is assured through reproducible calibration and testing of the extraction and
HPLC systems.
9.3 Interferences
9.3.1 Solvents, reagents, glassware, and other sample processing hardware may lead to
discrete artifacts and/or elevated baselines causing misinterpretation of
chromatograms.
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9.3.1.1 All materials used in the analysis shall be demonstrated to be free from
interferences by running method blanks initially and with each sample
batch (samples started through the extraction process at the same time, to a
maximum often). Specific selection of reagents and purification of
solvents by distillation in all-glass systems may be required.
9.3.1.2 Glassware and, where possible, reagents are cleaned by solvent rinse
and/or baking at 450°C for a minimum of 1 hour.
9.3.2 When used in conjunction with Method 1662, blanks extracted in that method are
treated as an integral part of this method.
9.3.3 Interferences co-extracted from samples may vary from source to source, depending
on the diversity of the site being sampled.
9.4 Safety
9.4.1 The toxicity or carcinogenicity of each compound or reagent used in this method has
not been precisely defined; however, each chemical should be treated as a potential
health hazard. Exposure to these chemicals must be reduced to the lowest possible
level.
9.4.2 The laboratory is responsible for maintaining a current awareness file of OSHA
regulations regarding the safe handling of the chemicals specified in this method. A
reference file of material safety data sheets (MSDSs) should also be made available to
all personnel involved in the chemical analysis. Additional information on laboratory
safety can be found in References 1 through 3.
9.4.3 Methylene chloride has been classified as a known health hazard. All steps in this
method which involve exposure to this compound shall be per-formed in an OSHA-
approved fume hood.
9.5 Apparatus and Materials
NOTE: Brand names, suppliers, and part numbers are for illustrative purposes only. No
endorsement is implied. Equivalent performance may be achieved using apparatus and materials
other than those specified here, but demonstration of equivalent performance meeting the
requirements of this method is the responsibility of the laboratory.
9.5.1 Equipment for glassware cleaning.
9.5.1.1 Laboratory sink with overhead fume hood.
9.5.1.2 Kiln: Capable of reaching 450°C within 2 hours and holding 450°C within
±10°C, with temperature controller and safety switch (Cress
Manufacturing Co, Sante Fe Springs, CA, B31H or X31TS, or equivalent).
9.5.2 Equipment for sample preparation.
9.5.2.1 Laboratory fume hood.
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9.5.2.2 Analytical balance: Capable of weighing 0.1 mg.
9.5.2.3 Glassware.
a. Disposable pipettes: Pasteur, 150 mm long by 5 mm i.d. (Fisher
Scientific 13-678-6A, or equivalent).
b. Glass pipettes: 1.0- and 10-mL, accurate to 1 % or better.
c. Volumetric flasks: Glass, 10- and 100-mL.
9.5.2.4 Sample vials: Amber glass, 2- to 5-mL with PTFE-lined screw-cap, to fit
FIPLC autosampler.
9.5.3 High-performance liquid chromatograph (FIPLC): An analytical system complete
with pumps, sample injector, column oven, and ultra-violet (UV) detector.
9.5.3.1 Pumping system: Capable of isocratic operation and producing a linear
gradient from 50% water/50% acetonitrile to 100% acetonitrile in 10
minutes (Waters 600E, or equivalent).
9.5.3.2 Sample injector: Capable of automated injection of up to 30 samples
(Waters 700, or equivalent).
9.5.3.3 Column oven: Capable of operation at room ambient to 50°C (Waters
TCM, or equivalent).
9.5.3.4 Column: Two Cig columns, 150 mm long by 4.6 mm i.d., 300 angstroms
(Vydac 201 TP5415, or equivalent) connected in series, preceded by one
Cig guard column, 30 mm long by 4.6 mm i.d., 300 angstroms (Vydac 201
GCC54T, or equivalent), operated at the conditions shown in Table 9-1.
9.5.3.5 Detector: UV operated at 254 nm (Waters 490E, or equivalent).
9.5.4 Data system.
9.5.4.1 Data acquisition: The data system shall collect and record LC peak areas
and retention times on magnetic media.
9.5.4.2 Calibration: The data system shall be used to calculate and maintain lists
of calibration factors (response divided by concentration) and multi-point
calibration curves. Computations of relative standard deviation
(coefficient of variation) are used to test calibration linearity.
9.5.4.3 Data processing: The data system shall be used to search, locate, identify,
and quantify the compounds of interest in each analysis. Displays of
chromatograms are required to verify results.
9.5.4.4 Statistics on initial (Section 9.8.2) and ongoing (Section 9.12.6)
performance shall be computed and maintained.
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9.6 Reagents
9.6.1 Solvents.
9.6.1.1 Sample preparation: Methylene chloride, distilled in glass (Burdick and
Jackson, or equivalent).
9.6.1.2 HPLC: Methanol, acetonitrile, and water, HPLC quality.
9.6.2 Standards: Purchased as solutions or mixtures with certification to their purity,
concentration, and authenticity, or prepared from materials of known purity and
composition. If compound purity is 96% or greater, the weight may be used without
correction to compute the concentration of the standard. If PAH in oil from drilling
mud is to be tested, the diesel oil standard used in this method should be from the oil
used on the drilling rig from which the mud sample is taken. If this oil is not
available, No. 2 diesel oil from a local source may be substituted.
9.6.2.1 Stock solutions: Prepare in methylene chloride or methanol and dilute In
acetonitrile for injection into the HPLC. Observe the safety precautions in
Section 9.4.
a. Diesel oil solutions
• Stock solution in methylene chloride (62.5 mg/mL): If QC extracts
from Method 1662 are to be tested, use the oil that was spiked in
that method. Weigh 6.25 g of diesel oil into a 100-mL ground-
glass-stoppered volumetric flask and fill to the mark with
methylene chloride.
• Diesel oil calibration solution (1.25 mg/mL): After the oil in the
stock solution (see bullet above) is completely dissolved, remove
1.00 mL and place in a 50-mL volumetric flask. Dilute to the mark
with acetonitrile. Mix thoroughly and transfer to a clean 150mL
bottle with PTFE-lined cap.
b. Polynuclear aromatic hydrocarbons-naphthalene, phenanthrene, and
indeno[l,2,3-cd]pyrene: Dissolve an appropriate amount of reference
material in a suitable solvent. For example, weigh 10.0 mg of
naphthalene in a 10-mL volumetric flask and fill to the mark with
methanol. After the naphthalene is completely dissolved, transfer the
solution to a 15-mL vial with PTFE-lined cap.
c. Stock solutions should be checked for signs of degradation prior to the
preparation of calibration or performance test standards.
9.6.2.2 PAH calibration standards (CAL): Dilute and mix the stock solutions
(Section 9.6.2.1b) in acetonitrile to produce the calibration standards
shown in Table 9-2. The three solutions permit the response of
phenanthrene to be measured as a function of concentration, and
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naphthalene and indeno[l,2,3-cd]pyrene permit the retention time window
for PAH to be defined. The medium-level solution is used for calibration
verification (Section 9.12.2).
9.6.2.3 Precision and recovery standard: The diesel oil calibration solution
(Section 9.6.2. la), second bullet) is used for initial precision and recovery
(IPR; Section 9.8.2) and ongoing precision and recovery (OPR, Section
9.12.6).
9.6.2.4 Stability of solutions.
a. When not being used, standards are stored in the dark at -20 to -10°C
in screw-capped vials with PTFE-lined lids. A mark is placed on the
vial at the level of the solution so that solvent loss by evaporation can
be detected. The vial is brought to room temperature prior to use. Any
precipitate is redissolved and solvent is added if solvent loss has
occurred.
b. Standard solutions used for quantitative purposes (Sections 9.6.2.1
through 9.6.2.3) shall be analyzed within 48 hours of preparation and
on a monthly basis thereafter for signs of degradation. Standards will
remain acceptable if the peak area remains within ±15% of the area
obtained in the initial analysis of the standard.
9.7 Calibration
9.7.1 Assemble the HPLC and establish the operating conditions in Table 9-2.
9.7.2 Retention time adjustment.
9.7.2.1 Inject 20 uL- of the medium level calibration standard (Table 9-2).
9.7.2.2 Locate the three peaks in this standard.
9.7.2.3 Adjust the initial solvent mixture, the isocratic hold, the gradient, and the
final isocratic hold until the retention times are within + 1 minute of the
retention times given in Table 9-2.
9.7.3 Minimum level: Analyze 20 uL of the low-level calibration standard (Table 9-2) and
verify that the HPLC instrument meets the minimum level for phenanthrene in Table
9-1.
9.7.4 External standard calibration.
9.7.4.1 Analyze 20 uL of each calibration standard (Table 9-2) beginning with the
lowest concentration and proceeding to the highest using to the procedure
in Section 9.11.
9.7.4.2 Record the areas for the phenanthrene peak and the height of the
phenanthrene peak in the high-level standard.
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9.7.4.3 Compute the ratio of response to amount injected (calibration factor) at
each concentration by dividing the area of the peak by the concentration of
the standard injected. Calculate the mean of the three values to produce an
average calibration factor.
9.7.4.4 Linearity: If the calibration factor is constant over the three point
calibration range (< 15% relative standard deviation), linearity through the
origin can be assumed; if not, the system shall be recalibrated.
9.7.5 The average calibration factor is verified on each working 8-hour shift by the
measurement of the medium-level calibration standard (Section 9.12.5).
9.7.6 Single-point calibration for diesel oil: Inject the precision and recovery standard
(Section 9.6.2.3) to produce a single calibration point for diesel oil.
9.7.6.1 Integrate the area from the retention time of naphthalene (including the
leading edge of the naphthalene peak) through the end of the indeno[l,2,3-
cd]pyrene peak or until the detector signal returns to a stable baseline,
whichever comes later, as shown in Figure 9-1.
9.7.6.2 Determine the calibration factor for diesel oil by dividing the integrated
area (9.7.6.1) by the diesel oil concentration (Section 9.6.2. la), second
bullet).
9.8 Quality Assurance/Quality Control
9.8.1 Each laboratory that uses this method is required to operate a formal quality
assurance program (Reference 4). The minimum requirements of this program consist
of an initial demonstration of laboratory capability, an ongoing analysis of standards
and blanks as a test of continued performance, analyses of spiked samples to assess
accuracy, and analysis of duplicates to assess precision. Laboratory performance is
compared to established performance criteria to determine if the results of analyses
meet the performance characteristics of the method. If the determination of PAH is to
be made on extracts from Method 1662, the quality control samples for initial
precision and recovery (IPR), spiked samples, duplicate samples, and ongoing
precision and recovery (OPR) samples from Method 1662 shall be substituted for
those in the QC tests below, and the specifications in Table 9-1 for extracts from
Method 1662 shall be met.
9.8.1.1 The analyst shall make an initial demonstration of the ability to generate
acceptable accuracy and precision with this method. This ability is
established as described in Section 9.8.2.
9.8.1.2 The analyst is permitted to modify this method to improve separations or
lower the costs of measurements, provided all performance requirements
are met. Each time a modification is made to the method, the analyst is
required to achieve the minimum level (Section 9.7.3) and to repeat the
procedure in Section 9.8.2 to demonstrate method performance.
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9.8.1.3 Analyses of spiked samples are required to demonstrate method accuracy
when extracts from Method 1662 are analyzed. The procedure and QC
criteria for spiking are described in Section 9.8.3.
9.8.1.4 Analyses of duplicate samples are required to demonstrate method
precision when extracts from Method 1662 are analyzed. The procedure
and QC criteria for duplicates are described in 9.8.4.
9.8.1.5 Analyses of blanks are required to demonstrate freedom from
contamination. The procedures and criteria for analysis of a blank are
described in Section 9.8.5.
9.8.1.6 The laboratory shall, on an ongoing basis, demonstrate through calibration
verification and analysis of the precision and recovery standard that the
analysis system is in control. These procedures are described in Sections
9.12.5 and 9.12.6.
9.8.1.7 The laboratory shall maintain records to define the quality of data that is
generated. Development of accuracy statements is described in Sections
9.8.3.2 and 9.12.6.4.
9.8.2 Initial precision and recovery (IPR): The initial precision and recovery test is
performed using the precision and recovery standard. If extracts from Method 1662
are to be analyzed, the extracts from the initial precision and recovery tests in that
method shall be used; otherwise, the laboratory shall generate acceptable precision
and recovery by performing the following operations.
9.8.2.1 Using diesel oil, prepare four separate aliquots of the precision and
recovery standard (Section 9.6.2.3). If extracts from Method 1662 are
analyzed, the extracts from the initial precision and recovery test in that
method shall be used. Analyze these aliquots using the procedure in
Section 9.11.
9.8.2.2 Using results of the set of four analyses, compute the average recovery (X)
of PAH in mg/mL and the standard deviation of the recovery (s) in mg/mL
for each aliquot by the external standard method (Sections 9.7.4 and
9.14.4).
9.8.2.3 Compare s and X with the corresponding limits for initial precision and
recovery in Table 9-1. If s and X meet the acceptance criteria, system
performance is acceptable and analysis of oil samples may begin. If,
however, s exceeds the precision limit or X falls outside the range for
accuracy, system performance is unacceptable. In this event, review this
method, correct the problem, and repeat the test.
9.8.3 Method accuracy: If extracts from Method 1662 are to be analyzed, the extract from
the accuracy test in that method shall be used; otherwise, an accuracy test is
unnecessary. The procedure for determining method accuracy is given in Section 8.3
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of Method 1662, and the specification for accuracy is given in Table 9-1 of this
method.
9.8.3. 1 Compare the percent recovery of PAH with the corresponding QC
acceptance criteria in Table 9-1. If the results of the spike fail the
acceptance criteria, and the recovery of the QC standard in the ongoing
precision and recovery test (Section 9. 12.6.3) is within the acceptance
criteria in Table 9-1, an interference may be present. In this case, the result
may not be reported for regulatory compliance purposes. If, however, the
results of both the spike and the ongoing precision and recovery test fail
the acceptance criteria, the analytical system is judged to be out of control
and the problem shall be identified and corrected, and the sample batch
reanalyzed.
9.8.3.2 As part of the QA program for the laboratory, method accuracy for
samples shall be assessed and records shall be maintained. After the
analysis of five spiked samples in which the recovery passes the test in
Section 9.8.3, compute the average percent recovery (P) and the standard
deviation of the percent recovery (P). Express the accuracy assessment as
a percent recovery interval from P - 2Sp to P + 2 Sp. For example, if P =
90% and Sp = 10% for five analyses of PAH in diesel oil, the accuracy
interval is expressed as 70 to 110%. Update the accuracy assessment on a
regular basis (e.g., after each five to ten new accuracy measurements).
9.8.4 Duplicates: If extracts from Method 1662 are to be analyzed, the extracts from the
duplicates test in that method shall be used. The procedure for preparing duplicates is
given in Section 8.4 of Method 1662, and the specification for RPD is given in Table
9-1 of this method. If extracts from Method 1662 are not to be analyzed, duplicates of
the precision and recovery standard (Section 9.6.2.3) are analyzed, and the
specification for RPD is given for PAH in diesel oil in Table 9-1 of this method.
9.8.4. 1 Analyze each of the duplicates per the procedure in Section 9.11 and
compute the results per Section 9. 14.
9.8.4.2 Calculate the relative percent difference (RPD) between the two results
per the following equation:
Dl~D2 I — 100 [9-1]
L J
D2)/2]
Where:
DI = Concentration of diesel oil in the sample
D2 = Concentration of diesel oil in the second (duplicate) sample
9.8.4.3 The relative percent difference for duplicates shall meet the acceptance
criteria in Table 9-1. If the criteria are not met, the analytical system is be
judged to be out of control, and the problem must be immediately
identified and corrected and the sample set re-extracted and reanalyzed.
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9.8.5 Blanks: If extracts from Method 1662 are to be analyzed, the extracts from blanks in
that method shall be analyzed in addition to the blanks in this method.
9.8.5.1 Rinse the glassware used in preparation of the extracts in this method with
acetonitrile and analyze a 20-uL aliquot of the rinsate using the procedure
in Section 9.11 and compute the results per Section 9.14.
9.8.5.2 If PAH is detected in a blank at greater than the method detection limit
(MDL) in Table 9-1, analysis of samples is halted until the source of
contamination is eliminated and a blank shows no evidence of
contamination.
9.8.6 The specifications contained in this method can be met if the apparatus used is
calibrated properly, and then maintained in a calibrated state. The standards used for
initial precision and recovery (TPR, Section 9.8.2) and ongoing precision and recovery
(OPR, Section 9.12.6) should be identical, so that the most precise results will be
obtained. The HPLC instrument will provide the most reproducible results if
dedicated to the settings and conditions required for the analyses given in this
method.
9.8.7 Depending on specific program requirements, field replicates and field spikes of
diesel oil into samples may be required when Method 1662 and this method are used
to assess the precision and accuracy of the sampling and sample transportation
techniques.
9.9 Sample Collection, Preservation, and Handling
9.9.1 Oil samples are collected in 20- to 40-mL vials with PTFE- or aluminum-foil-lined
caps and stored in the dark at -20 to -10°C.
9.9.2 If extracts from Method 1662 are to be analyzed, the laboratory should be aware that
sample and extract holding times for this method have not yet been established.
However, based on tests of wastewater for the analytes determined in this method,
samples shall be extracted within 7 days of collection and extracts shall be analyzed
within 40 days of extraction.
9.9.3 As a precaution against analyte and solvent loss or degradation, sample extracts are
stored in glass bottles with PTFE-lined caps, in the dark, at -20 to -10°C.
9.10 Dilution of Oil and Extracts
9.10.1 Neat oil samples: If oil is received in neat form, it should be diluted to bring the
concentration within the range of the instrument. If the oil is No. 2 diesel oil, the
appropriate concentration will be approximately 2,000 ug/mL. Mineral oils and other
oils containing a lesser PAH content will require less dilution.
9.10.2 Extracts from Method 1662: If extracts of samples from Method 1662 are to be
analyzed, these extracts (from Section 10.4.2 of that method) are analyzed undiluted
unless diesel oil is known or suspected to be present. Extracts of QC samples (TPR,
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OPR, matrix spikes, and duplicates) from Method 1662 are diluted by a factor of 10
to bring them within the range of the HPLC.
9.10.3 Dilution of neat oil expected to be diesel oil.
9.10.3.1 Weigh 100 mg into a 10-mL volumetric flask and dilute to the mark with
methylene chloride to produce a concentration of 10 mg/mL. Stopper and
mix thoroughly.
9.10.3.2 Using a calibrated 1.0-mL volumetric pipette, withdraw 1.0 mL of the
solution and place in a 10-mL volumetric flask. Then withdraw an
additional 0.25 mL of the solution and place in the 10-mL volumetric flask
(for a total of 1.25 mL). Fill to the mark with acetonitrile to produce a
concentration of 1.25 mg/mL (1250 ug/mL). This solution will be near,
but not above, the limit of the calibration range and will match the
concentration of the QC samples from Method 1662 (assuming 100%
recovery).
9.11 High-Performance Liquid Chromatography
9.11.1 Table 9-2 summarizes the recommended operating conditions for the HPLC. Included
in this table and in Table 9-1 are retention times and the minimum level that can be
achieved under these conditions. An example of the separation achieved for diesel oil
by the multiple HPLC column system is shown in Figure 9-1. Other HPLC columns,
chromatographic conditions, or detectors may be used if the requirements for the
minimum level (Section 9.7.3) and initial precision and recovery (Section 9.8.2) are
met.
9.11.2 Calibrate the system as described in Section 9.7 or verify calibration as described in
Section 9.12.
9.11.3 Analysis of extracts.
9.11.3.1 Inject 20 uL of the sample extract, Method 1662 extract, or diluted QC
extract into the HPLC using a high-pressure syringe or a constant-volume
sample-injection loop. Record the volume injected to the nearest 0.1 uL.
9.11.3.2 Upon injection, begin the solvent program used in calibrating the column
(Section 9.7.2.3). Record the signal from the time of injection until the
detector returns to a stable baseline. Return the solvent to the initial
conditions.
9.11.3.3 Using the retention-time data determined during calibration, integrate the
area from the retention time of naphthalene (including the leading edge of
the naphthalene peak) through the end of the indeno[l,2,3-cJ]pyrene peak
or until the detector signal returns to a stable baseline, whichever comes
later.
9-10
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9.11.4 If the height of the response during the period recorded (Section 9.11.3.2) exceeds the
height of the response for phenanthrene during calibration (Section 9.7.4.2), dilute the
extract by successive factors of 10 with acetonitrile and reanalyze until the response
is within the calibration range.
9.12 HPLC System and Laboratory Performance
9.12.1 At the beginning of each 8-hour shift during which analyses are performed, HPLC
calibration and system performance are verified. For these tests, analysis of the
medium level calibration standard (Table 9-2) and of the diluted extract of the
precision and recovery standard (Section 9.6.2.3) shall be used to verify all
performance criteria. Adjustment and/or recalibration (per Section 9.7) shall be
performed until all performance criteria are met. Only after all performance criteria
are met may samples and blanks be analyzed.
9.12.2 Inject 20 uL of the medium-level calibration standard (Table 9-2) into the HPLC
instrument according to the procedure in Section 9.11.
9.12.3 Retention time: The absolute retention times of the naphthalene, phenanthrene, and
indeno[l,2,3-cd]pyrene peaks shall be within ±30 seconds of the respective retention
times in the initial calibration (Section 9.7.2.3).
9.12.4 HPLC resolution: Resolution is acceptable if the peak width at half-height of the
phenanthrene peak is less than 30 seconds.
9.12.5 Calibration verification: Compute the concentration of phenanthrene based on the
average calibration factor (Section 9.7.4.4). The concentration shall be within the
limits in Table 9-1. If calibration is verified, system performance is acceptable and
analysis of blanks and QC samples may begin. If, however, the concentration falls
outside of the calibration verification range, system performance is unacceptable. In
this case, correct the problem and repeat the test, or recalibrate (Section 9.1 A).
9.12.6 Ongoing precision and recovery (OPR): If the extract is from Method 1662, the OPR
standard from that method shall be used and the specification for the OPR from
Method 1662 in Table 9-1 shall be met; if not, a sample of diesel oil shall be diluted
per the procedure in Section 9.10 and shall be used for the OPR test.
9.12.6.1 Analyze the appropriate OPR standard.
9.12.6.2 Compute the concentration of PAH in this standard per Section 9.14.
9.12.6.3 Compare the concentration with the limits for ongoing precision and
recovery in Table 9-1. If the concentration is in the range specified, the
analytical processes are in control and analysis of blanks and samples may
proceed. If, however, the concentration is not in the specified range, these
processes are not in control. In this event, correct the problem, re-extract
the sample batch if the OPR is from Method 1662, or redilute the oil
sample (per Section 9.10.3). and repeat the ongoing precision and
recovery test.
-------
9.12.6.4 Add results which pass the specification in Section 9.12.6.3 to initial and
previous ongoing data. Update QC charts to form a graphic representation
of continued laboratory performance. Develop a statement of laboratory
data quality for each analyte by calculating the average percent recovery
(R) and the standard deviation of percent recovery (Sr). Express the
accuracy as a recovery interval from R - 2Sr, to R + 2 Sr. For example, if
R = 95% and Sr = 5%, the accuracy is 85 to 105%.
9.13 Qualitative Identification
9.13.1 Qualitative determination is accomplished by comparison of data from analysis of a
sample or blank with data from analysis of the calibration verification standard
(Section 9.12.5).
9.13.2 PAH is identified in the sample by the presence of peaks and/or an elevated baseline
(hump) between the retention times of the naphthalene and indeno[l,2,3-cd]pyrene
peaks (Section 9.11.3.3), as shown in Figure 9-1. The experience of the analyst shall
weigh heavily in interpretation of the chromatogram.
9.14 Quantitative Determination
9.14.1 Using the data system, compute the concentration of the PAH detected in the solution
injected into the HPLC (in ug/mL) using the calibration factor (Section 9.7.4).
9.14.2 Concentration of PAH in oil: If neat oil was analyzed, the concentration of PAH in
the oil is determined using the following equation:
c«= w
Where:
C0 = Concentration of PAH in the oil sample
Cp = Concentration of PAH measured (from Sections 9.11.4 and 9.14.1)
C; = Concentration of oil in the solution injected into the HPLC (from
Sections 9.10.3.2, 9.11.4, and 9.14.1)
9.14.3 Concentration of diesel oil in QC extracts from Method 1662: Calculate the
concentration of diesel oil in QC extracts from Method 1662 by integrating the area
per Section 9.7.6.1 and using the calibration from Section 9.7.6.2 of this method,
taking into account the dilution of these extracts (Section 9.10.2).
9.14.4 Concentration of PAH in oil from Method 1662: The PAH content of oil is
complicated by the splitting and possible dilution of these extracts.
9.14.4.1 Concentration in undiluted extracts: This concentration is determined by
Equation 9-3:
C = VexCP = 5XICP r931
1/5 xWT WT L J
-------
Where:
C0 = Concentration of PAH in the oil sample
Ve = Amount of extract split for HPLC analysis, in mL (1.0 mL)
Cp = Concentration of PAH measured
WT = Weight of oil in the concentration tube in Method 1662 (Section
11.5.5 of Method 1662)
1/5 = Fraction of this weight used for the PAH determination
9.14.4.2 Concentration in diluted extracts: If the extract was diluted by a factor of
10 (Section 9.10.3 or 9.11.4), the concentration determined in Section
9.14.4.1 is multiplied by 10.
9.14.5 If the concentration is to be expressed as weight percent, C0 is multiplied by 0.1.
9.14.6 Report results to three significant figures without correction for recovery.
9.15 Method Performance
This method was validated in a single laboratory (Reference 6) using samples of hot-rolled
drilling mud (Reference 7).
9.16 References
1. "Carcinogens-Working With Carcinogens." Department of Health, Education, and
Welfare, Public Health Service, Centers for Disease Control [available through National
Technical Information System, 5285 Port Royal Road, Springfield, VA 22161, document
no. PB2772561: August 1977.
2. "OSHA Safety and Health Standards, General Industry [29 CFR 1910], Revised."
Occupational Safety and Health Administration, OSHA 2206. Washington, DC: January
1976.
3. "Safety in Academic Chemistry Laboratories (3rd Edition)." American Chemical Society
Publication, Committee on Chemical Safety. Washington, DC: 1979.
4. "Handbook of Analytical Quality Control in Water and Wastewater Laboratories."
USEPA, EMSL-Ci, EPA-600/4-79-019. Cincinnati, OH: March 1979.
5. " Standard Practice for Sampling Water," ASTMAnnual Book of Standards, Part 31,
D337076, ASTM. Philadelphia, PA: 1980.
6. "Determination of Polynuclear Aromatic Hydrocarbons and Diesel by Modified EPA
Method 9310." Prepared for the American Petroleum Institute c/o Shell Development Co,
Westhollow Research Center, 3333 Highway 6 South, Houston, TX 77082 by Analytical
Technologies Inc., 225 Commerce Drive, Fort Collins, CO 80524: March 29 1991,
April 12, 1991, and August 18, 1992.
7. "Results of the API Study of Extraction and Analysis Procedures for the Determination
of Diesel Oil in Drilling Muds (Final Report)." American Petroleum Institute, Offshore
-------
Effluent Guidelines Steering Committee, Technology Work Group, Prepared by 1C.
Raia, Shell Development Co. Houston, TX: April 18, 1991.
Table 9-1. Performance Data and Method Acceptance Criteria for PAH
Criterion
Minimum level °
Method Detection Limit d
Units
ug/mL
ug/mg
PAH in Diesel Oil a
100
7.6
Diesel Oil in Mud
Extract b
—
—
Phenanthrene
0.1
—
Initial prec and recov
Precision (std dev)
PAH in diesel oil e
Diesel in mud extract f
Recovery
PAH in diesel oil e
Diesel in mud extract f
Calibration verification 8
mg/mL
mg/mL
mg/mL
mg/mL
Ug/mL
120
—
1,090-1,340
—
—
—
0.55
—
0.84-1.95
—
—
—
—
—
0.39-0.61
Ongoing prec and recov
PAH in diesel oil e
Diesel in mud extract f
Matrix spike recovery f
Duplicates
mg/mL
mg/mL
pet
RPD
1,010-1,450
—
—
9.5
—
0.76-2.15
0.43-2.39
44
—
—
—
—
a CAS Registry number 68534-30-5; No. 2 diesel oil used for these tests.
b From Method 1662.
0 This is a minimum level at which the analytical system shall give recognizable signals and acceptable calibration
points..
d 40 CFR Part 136, Appendix B; MDL is measured as PAH in oil.
e Test concentration of diesel oil = 1,250 ug/mL.
f Test concentration in diluted extract = 1.25 mg/mL.
8 Test concentration = 0.50 ug/mL.
Table 9-2. HPLC Calibration Data
Analyte
Naphthalene
Phenanthrene
Indeno [123 -«/]pyrene
Diesel oil b
Retention Time a
(minutes)
7.6
10.3
18.9
7.4-20.0
Calibration Solution Concentration (jig/mL)
Low
—
0.1
—
100
Medium
5
0.5
0.5
400
High
—
2.0
—
2,000
a Column system: Two Cis columns (150 mm long by 4.6 mm i.d., 300 angstroms) connected in series,
preceded by one Cis guard column (30 mm long by 4.6 mm i.d., 300 angstroms). Column temperature 30°C;
solvent flow rate 1.5 mL/min; linear gradient from 50% water/50% acetonitrile at injection to 100%
acetonitrile in 10 minutes, hold at 100% acetonitrile for 15 minutes.
b Diesel oil is calibrated separately using a single point calibration (Section 9.7.6).
9-14
-------
00
05
CD
0.
CO
Three-Component Standard
i
£
CO
CD
Q_
CD
I
Q.
"O"
u
CO_
O> CM
05 ,-
S2 "o*
I CD
c
No. 2 Diesel Oil
0.00
0.50
1.00 1.50
x 101 minutes
2.00
Figure 9-1. Liquid Chromatography of the Three-Component Standard and of No. 2 Diesel
Oil
9-15
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10. METHOD FOR CONDUCTING A SEDIMENT TOXICITY TEST WITH LEPTOCHEIRUS
PLUMULOSUS AND NON-AQUEOUS DRILLING FLUIDS OR SYNTHETIC-BASED DRILLING
MUDS (EPA METHOD 1644)
10.1 Summary of Method
This test method describes the procedures for obtaining data regarding the effects of non-
aqueous drilling fluids (NAFs) or synthetic based drilling muds (SBMs) on the marine
amphipod, Leptocheirusplumulosus. EPA is regulating the sediment toxicity in NAFs
and SBMs that are discharged from oil and gas extraction facilities in coastal and
offshore waters as an indicator for toxic pollutants (see 40 CFR 435.13, 435.15, 435.43,
and 435.45). EPA established the use of this test and related limits to encourage the use
of less toxic drilling fluids and additives. The sediment toxicity of the NAF-cuttings at
the point of discharge is measured by this modified sediment toxicity test using a natural
sediment and Leptocheirus plumulosus as the test organism.
EPA promulgated the use of this method with revisions to the Oil and Gas Extraction
effluent guidelines (U.S. EPA, 2001). This initial method was consistent with ASTM
Standard Guide E 1367-92 (ASTM 1997). Subsequent to this rulemaking EPA updated
this method in the NPDES permit for the Western Gulf of Mexico (GMG290000,
Appendix A) to be consistent with ASTM E 1367-99 (ASTM 2000). This method is a re-
publication of the sediment toxicity test in NPDES permit for the Western Gulf of
Mexico (GMG290000, Appendix A).
10.2 Test Requirements and Materials
10.2.1 Test Species (Leptocheirusplumulosus)
10.2.1.1 Description: Leptocheirus plumulosus is an infaunal amphipod that is
indigenous to subtidal regions along the east coast of the U.S. This
amphipod constructs U-shaped burrows in the top 5 cm of fine sand to
silty clay sediments (ASTM E 1367-99). As a result of its broad salinity
and particle size tolerances, it is a desirable test species for a variety of
toxicity testing programs.
10.2.1.2 Collection and Handling: In the field, amphipods can be collected using
sediment grab samplers such as Peterson and Ponar dredges. This species
has been collected in various tributaries of the Chesapeake Bay for various
toxicity testing programs (ASTM E 1367-99). The contents of each grab
should be sieved through a 500 mesh screen. The sediment and organisms
retained on the screen are gently rinsed into plastic buckets containing
sediment and water from the collection site. These buckets are quickly
transported back to the laboratory and aerated. See ASTM E 1367-99 for
more details on collection and handling.
10.2.1.3 Holding and Acclimation: Amphipods can be placed in aquaria containing
a 1-2 cm deep layer of collection site sediment that has been sieved
through a 500 mesh screen. Amphipod density should be about 200-300
per 40 L aquarium with vigorous aeration. Two to three days are sufficient
KM
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for acclimation to test conditions, and during this period a gradual change
over from site water to test water is recommended (ASTM E 1367-99).
10.2.1.4 Environmental Tolerances: L. plumulosus is tolerant of a broad salinity
range, from near 0 to 33 g/kg (%o) (ASTM E 1367- 99). This species has
demonstrated up to 100% survival in >90% silt-clay sediment and an
average of 85% survival in >95% sand/gravel sediment (ASTM E 1367-
99). The ASTM data are consistent with data published from other studies
indicating that L. plumulosus is tolerant of sandy and silty sediments. For
example, Schlekat et al. (1992) noted a mean survival of 97.5% whenZ.
plumulosus was exposed for 10 days to field collected sediments ranging
from 98.1% sand to 96.5% fines. Further, this species was collected in the
field in sediments consisting of 99.9% sand and 92.1% fines, indicating
that L. plumulosus is a generalist and can thrive in a variety of sediment
types (Schlekat et al. 1992).
However, the fine fraction of sediments in the Schlekat et al. study did not
exceed 55% clay, indicating that the fine fraction was a mixture of silt and
clay sized particles. Data from other studies indicated that this species is
intolerant of sediments high in clay content. McGee et al. (1999) noted
acceptable survival when this species was exposed to Baltimore Harbor
sediments containing up to 72% clay. However, Emery et al. (1997) noted
significantly reduced amphipod survival when L. plumulosus was exposed
for 10 days to Magothy River, Maryland sediment (amended with beach
sand and kaolinite clay) containing 84%, 90%, and 100% clay.
These data indicated that the tolerance range of this amphipod to clay
content is between about 72 to 84%. As such, caution should be used
when conducting L. plumulosus toxicity tests with sediments with clay
content greater than about 70%. This should not have a significant impact
on using this species in the NAF and SBM toxicity testing program, since
field sediments seldom exceed 70% clay content (Suedel and Rodgers,
1991).
10.2.2 Culture Methods (Leptocheirusplumulosus)
10.2.2.1 Overview: Populations of L. plumulosus can be maintained through
several generations in the laboratory. The culture conditions specified in
ASTM E 1367-92 and E 1367-99 are provided in Table 1.
10-2
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Table 10-1. Culture conditions for L. pluniulosus. Conditions listed are consistent with
culture conditions specified in ASTM E 1367-92 and subsequent updates (E 1367-99)
Parameter
Temperature
Salinity
Light Quality
Illuminance
Photoperiod
Culture Chamber
Sediment Volume
Renewal of overlying water
Number of
organi sm s/chamb er
Feeding
Aeration
Overlying Water
Overlying water quality
Conditions
20±1 °C
20±l%o
Wide-spectrum fluorescent or cool white lights
500-1,000 lux
14h light: 1 Oh dark
Shallow plastic tubs or glass aquaria
1-2 cm depth at bottom of each culture chamber
Static renewal (30-50% water volume change 2-4
times per week)
Start with about 300 mixed age (mostly immature and
young adults) individuals per chamber
0. 1 to 0.5 g dry mixture 2-3 times per week (see section
10.2.2.2)
Continuous gentle to moderate aeration so as to not
suspend sediments
Clean natural or synthetic seawater
Salinity, temperature, and ammonia during culture
start-up
In addition to the conditions provided in Table 1, there are other
conditions that are important in maintaining healthy L. plumulosus
cultures, including identifying a source of clean sediment, sieving
sediments before use, and the quality of the raw materials used to prepare
their food. Preferably, the sediment and water used to culture the
amphipods should be collected from the same area as those used in NAF
tests. Fine-grained sediments have been shown to be suitable for this
purpose (ASTM E 1367-92). Sediments collected in the field for culturing
purposes should be first sieved through a 2,000 m mesh sieve to remove
large debris and then through a 500 m mesh sieve to remove any
indigenous organisms. L. plumulosus cultures should be maintained at
20±1 °C and 20±l%o salinity. If used, natural seawater should be filtered
through a 5 micron filter before adding to cultures. New culture chambers
should be aerated and allowed to equilibrate overnight before adding
amphipods. Water used to start a new culture chamber should be renewed
24 h after initiation and before amphipods are added to culture chambers;
otherwise, culture water should be renewed in conjunction with feeding.
Cultures should be observed daily to ensure sufficient aeration. An
abundance of amphipods on the sediment surface during daylight hours
may indicate insufficient dissolved oxygen or overcrowding, as
amphipods typically remain in their burrows unless they are searching for
food or a mate. Culture chambers should be terminated and restarted with
fresh sediment about once every 8 weeks to avoid overcrowding.
10-2
-------
Overcrowding may lead to stress due to food or space limitations, and may
also result in reduced female fecundity, thus reducing the relative health of
the population of amphipods in a given culture chamber.
Cultures should be routinely inspected for the presence of indigenous
worms and copepods, a microbial build-up, or black and sulfurous
conditions beneath the sediment surface. Microbial growth appears as a
white or gray growth associated with uneaten food, and is indicative of
overfeeding. Presence of indigenous species, excess microbial growth, or
black and sulfurous conditions may necessitate discarding the affected
culture chamber.
10.2.2.2 Feeding: A mixture of micro-algae, yeast, fish food flakes, alfalfa powder,
ground cereal leaves, and shrimp maturation feed has been used to feed
cultures (ASTM E 1367-92 and E 1367-99). Micro-algae used in culturing
include Pseudoisochrysis paradoxa, Phaeodactylum tricornutum, and
Tetraselmis suecica mixed in equal parts on a volume basis. These algae
provide a source of fatty acids that may otherwise be absent in the diet. In
practice, however, it should be noted that L. plumulosus has been cultured
successfully without the algal mixture and the yeast. The dry food portion
of the diet that has been used to successfully culture L. plumulosus is
shown in Table 2.
Table 10-2. Dry food portion of the diet that has been used to successfully culture L.
plumulosus
Dietary Component
Fish food flakes (e.g., TetraMin®)
Alfalfa powder
Ground cereal leaves (dried wheat leaves)
Shrimp maturation feed (e.g., Neo-Novum®)
Proportion
48.0%
24%
24%
4.0%
This dry food mixture should be homogenized into a fine powder and fed
to each culture chamber at a rate of 0.1 to 0.5 g two to three times per
week, depending on culture densities. Overfeeding may result in microbial
build-up on the sediment surface. The quality of the alfalfa powder and
dried wheat leaves may not be consistent among suppliers, thus potentially
adversely affecting culture performance. Feeding should occur
immediately after culture water changes.
10.2.3 Obtaining Amphipods for Starting a Test (Leptocheirusplumulosus)
10.2.3.1 Immature and adult amphipods of mixed sexes and approximately 3 to 5
mm in length (as measured from the base of the first antenna to the end of
the third pleon segment along the dorsal surface) are used in toxicity tests,
as they are easier to handle and count than younger individuals. Gravid
females are not used in testing.
10-4
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10.2.3.2 The 3 to 5 mm size class individuals are passed through a 1,000 m mesh
sieve and are retained on a 710 m mesh sieve. A 500 m mesh sieve has
been used previously to retain amphipods of the size needed, but this will
result in a wider size range of amphipods used for testing.
10.2.3.3 In preliminary NAF experiments, this wide size range may have
contributed to variability in mortality observed that was not present when
the 710 m mesh sieve was used to retain amphipods in later experiments.
The amphipods passing through a 1000 m mesh sieve but trapped on a 710
m mesh sieve provide a more uniform size range of animals that is thought
to decrease the previously-observed variability in mortality. Laboratories
are encouraged to use this type of approach to reduce the variability in the
size of amphipods used in the NAF/SBM testing program.
10.2.4 Control Sediments
10.2.4.1 Overview: Control sediment must meet certain minimum requirements to
be used in the NAF/SBM testing program. The primary requirement is that
the sediment should be able to support L. plumulosus in cultures for
extended periods of time. This will ensure that the sediment is chemically
nontoxic and that the physical and chemical characteristics of the sediment
(e.g., total organic carbon, particle size distribution, and moisture content)
are within the tolerance range of the test species. It is expected that
separate aliquots of the culture sediment will also be used as a control
sediment to be amended by NAFs or SBMs in the NAF/SBM testing
program. Any modifications made to the control sediments should be
noted in the report.
10.2.4.2 Characterization: Sediments used in testing should be characterized for
total organic carbon (TOC), particle size distribution (sand, silt, and clay),
and percent water content. These parameters have been shown to influence
the results of NAF/SBM toxicity to L. plumulosus in initial experiments.
Variations in these sediment characteristics should be quantified so that
potential effects of these parameters on test results can be closely
monitored.
10.2.4.3 Collection: Control sediments should be collected from the amphipod
collection site or from another area that can provide a consistent source of
sediment with characteristics within the tolerance range of L. plumulosus.
Sediments showing evidence of chemical contamination should not be
used in the NAF/SBM testing program. Any site water overlying the
sediment should be retained so that fine particles suspended in the water
can be re-combined with the sediment before use. Sediment salinity and
temperature should be recorded at the time of collection. Sediment
collected for use should be homogenized and a composite sample prepared
for analysis for the parameters outlined above.
10.2.4.4 Sieving: Sediments collected in the field for culturing and testing purposes
should be first press-sieved through a 2,000 or similar mesh sieve to
-------
remove large debris and then through a 500 mesh sieve to remove any
indigenous organisms. Sediments have also been press-sieved through a
250 to 350 mesh sieve prior to testing to aid in the enumeration of
amphipods on a 500 mesh sieve at test termination.
10.2.4.5 Storage: The control sediment should be stored in plastic or glass
containers at 4 °C until test initiation. The sediment should be stored in the
dark and should not be allowed to freeze or dry out during storage (ASTM
E 1367-92).
10.2.5 Test Water
10.2.5.1 Water used in the NAF/SBM testing program should be available in
sufficient quantities and be acceptable to L. plumulosus. The minimum
requirement for acceptable water for use in the NAF/SBM testing program
is that healthy test organisms survive in the water, and in the water plus
control sediment, for the duration of holding and testing without showing
signs of disease or stress (ASTM E 1367-99). Another test for
acceptability of the test water would be its successful use in the culturing
of L. plumulosus (with the control sediment).
10.2.5.2 Natural seawater or synthetic salt water can be used in the NAF/SBM
testing program. Natural salt water should be obtained from an
uncontaminated area known to support a healthy, reproducing population
of L. plumulosus or similar sensitive species. Reconstituted salt water can
be prepared by adding commercially available sea salt in specified
quantities. Natural seawater should be filtered by passing through a 5
micron filter before use. The reader is referred to ASTM E 1367-92 or E
1367-99 for more information concerning test water.
10.3 Procedures
10.3.1 Mixing NAFs or SBMs with Control Sediment
EPA Method 1646 describes the method that should be used for amending control
sediments with NAFs and SBMs. The control sediment should be sieved and
homogenized before wet to dry weight ratio and density determinations are made and
before NAFs are added to the control sediment. The following steps were given in
EPA Method 1646 for mixing NAFs and SBMs with control sediments (parentheses
were added here to provide additional information).
10.3.1.1 Determine the wet to dry weight ratio for the control sediment (three
replicates of 30 g each has been used successfully)
10.3.1.2 Determine the density (g/ml) of the control sediment (three replicates of
>25 ml is suitable for this purpose);
10.3.1.3 Determine the amount of NAF or SBM needed to obtain a desired test
concentration;
-------
10.3.1.4 Determine the amount of wet sediment required;
10.3.1.5 Determine the amount of dry sediment in kilograms for each test
concentration;
10.3.1.6 Determine the amount of NAF or SBM required to amend the control
sediment at each test concentration;
10.3.1.7 Mix NAF or SBM with control sediment;
10.3.1.8 Test for homogeneity of NAF or SBM in sediment, and;
10.3.1.9 Mix sufficient quantities of NAF or SBM with control sediment for each
treatment of amended or spiked sediment.
10.3.2 Estimate of Require NAF and SBM Required
The six steps given above for NAFs can also be applied to SBMs, except that the
third bullet in Step 3 requires a measurement of the density of the SBM. The density
of the SBM can then be used to estimate the quantity required for the desired test
concentration. Refer to the formulas below for NAF and SBM calculations:
WAT7Upm,irpHrcrt _ [Cone. Desired (mg/kg)] [Dry Weight Sediment (g)]
NAF Required (g) - iQOOg/kg lOOOmg/g [10-1]
SBM Required (g) = [Cone. Desired (ml/kg)] x pry Weight Sediment (kg) x [SBM Density (g/ml)] [ 10-2]
See 40 CFR 435.13, 435.15, 435.43, and 435.45 for more information on this procedure.
10.3.3 Mixing Procedure
Mixing the NAF or SBM with the control sediment should be accomplished by
following these steps. The control sediment alone should also be subjected to the
mixing procedure to ensure mixing has no effect on sediment toxicity.
10.3.3.1 Place appropriate amounts of weighed NAF or SBM into a stainless steel
mixing bowl.
10.3.3.2 Tare the mixing bowl weight.
10.3.3.3 Add appropriate amount of control sediment.
10.3.3.4 Mix for 9 to!5 minutes with a hand-held mixer equipped with stainless
steel blades (e.g., KitchenAid Model KHM6).
10.3.3.5 As appropriate, test mixing homogeneity as described in Section 10.3.4.
10.3.4 Homogeneity of Mixing
-------
10.3.4.1 Tests for homogeneity of mixing should be performed, preferably in the
procedure development phase (see 40 CFR435.13, 435.15, 435.43, and
435.45) by each laboratory performing NAF/SBM toxicity testing. This is
to ensure that the NAF or SBM, which can be difficult to homogenize with
control sediments, can be evenly mixed with the control sediment by each
testing laboratory.
10.3.4.2 EPA Method 1646 specifies that the coefficient of variation (CV) for a
minimum of three replicate samples of the NAF/control sediment mixture
must be less than 20%. Determinations of C V should be based on total
petroleum hydrocarbon (TPH) content of the NAF or SBM as measured
by EPA Methods 3550A and 8015M. If the initial CV is 20%, then the
NAF/SBM-sediment mixture must be re-mixed and reanalyzed until the
20% CV limit is achieved.
10.3.4.3 Homogeneity measurements should be made on the lowest and highest
NAF concentrations for a given test. Laboratories should validate mixing
efficiency via TPH measurements (as outlined above) of the low and high
NAF concentrations. The homogeneity measurements should be made at
least once per year.
10.4 Recommended Test Conditions
10.4.1 The recommended test conditions for conducting the 10-day or 96-hr sediment
toxicity test with L. plumulosus are summarized in Table 3 and are consistent with
methods presented in ASTM E 1367-92 and subsequent updates (E 1367-99). Tests
should be conducted at 20±1 °C and 20±1%0 salinity with a 14h light; 10 h dark
photoperiod at approximately 500-1,000 lux (or about 46 to 93 footcandles). Test
chambers are 1-L glass containers with about a 10 cm inside diameter opening (or
similar glass containers) that can contain about 150 ml sediment and 600 ml
overlying water to achieve a 4:1 (v/v) water to sediment ratio.
10.4.2 There are five (5) test concentrations plus a control for each NAF and SBM test. Five
(5) replicates are included for the control sediment (E 1367-99) and for each test
concentration.
10.4.3 The control sediment/test material mixture and test water should be added to test
chambers the day before amphipods are added. This will allow for suspended
particles to settle and allow time for equilibration of temperature and the sediment-
water interface.
10.4.4 After the overnight equilibration period, amphipods are randomly distributed to each
test chamber. Twenty amphipods are added to each replicate and there are five
replicates per test treatment.
10.4.5 Amphipods caught on the water surface can be pushed under with a glass rod.
Individuals that have not burrowed within 5 to 10 minutes can be replaced, unless
they are exhibiting an avoidance response. Amphipods are not removed at any time
during the course of the toxicity test even if they appear dead. Test water is not
-------
renewed (i.e., static) and the amphipods are not fed during the exposure period. The
toxicity test is terminated after 96 hours or 10 days for SBMs and NAFs respectively.
10.4.6 Temperature, salinity, pH, and dissolved oxygen (DO) should be monitored daily.
10.4.7 Ammonia should also be monitored in overlying water to ensure that the
concentrations of this constituent do not exceed the tolerance range of the test species.
For L. plumulosus, this is about 60 mg/L (as total ammonia) at pH 7.7 in 10-day tests
(U.S. EPA, 1994). Ammonia has not been a problem in initial L. plumulosus 96-hr
and 10-day tests with various NAFs.
Table 10-3. Conditions for conducting 96-hour NAF and 10-day SBM sediment toxicity
tests with L. plumulosus. Conditions listed are consistent with test conditions specified in
ASTM E 1367- 92 and subsequent updates (E 1367-99) unless otherwise noted
Parameter
Test Type
Temperature
Salinity
Light Quality
Illuminance
Photoperiod
Test Chamber
Sediment Volume
Overlying water volume
Renewal of overlying water
Size and life stage of
amphipods
Number of
organi sm s/chamb er
Number of test concentrations
Number of replicate
chambers/treatment
Feeding
Aeration
Overlying Water
Overlying water quality
Test duration
Endpoint
Test acceptability
Conditions
Static whole sediment toxicity test
20±1 °C
20±l%o
Wide-spectrum fluorescent or cool white lights
500-1,000 lux
14h light: 1 Oh dark*
1-L glass beaker or jar
150 ml (2 cm depth)
600 ml (4: 1 [v/v] water to sediment ratio)
None
3-5 mm; immature and adult
20
5
5 in both controls and test treatments
None
Water in each test chamber should be
aerated throughout the test.
Clean natural or synthetic seawater
Temperature, salinity, pH, andD.O.
daily; ammonia, as needed
96 hours or 10 days
Survival
Minimum mean control survival of 90% and satisfaction of
criteria outlined in Table 4.
*Note: Although ASTM E 1367 specifies 16h light:8h dark, the photoperiod was changed to 14h light: lOh dark to
be consistent with the Mysidopsis bahia bioassay for drilling fluids (EPA Method 1619).
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10.5 Biological Data
10.5.1 Mortality is the endpoint for L. plumulosus at the end of the exposure period. At test
termination, the contents of each test chamber (amphipods plus test sediment) are
sieved through a 500 mesh screen to remove amphipods.
10.5.2 Material retained on the screen should be rinsed into a sorting tray with clean salt
water.
10.5.3 The total numbers of live and dead amphipods should be recorded.
10.5.4 Missing animals are presumed to have died and decomposed during the test and
disintegrated.
10.5.5 Amphipods should be counted alive if there are any signs of movement, such as a
neuromuscular pleopod twitch (ASTM E 1367-99). Gentle prodding may be used to
elicit movement.
10.6 Quality Control
10.6.1 Table 4 provides the acceptability requirements for the 10-day NAF and 96-hr SBM
test. The primary acceptability requirement for NAF testing is as follows.
10.6.2 A toxicity test is unacceptable if more than a total of 10% of the control organisms
die, or if the coefficient of variation (CV) of control survival is equal to or greater
than 40%.
10.6.3 If this acceptability requirement is not met, then the data should be discarded and the
experiment repeated.
10.6.4 If this requirement is met, then the other acceptability requirements in Table 4 should
be reviewed and a determination made as to the acceptability of the data.
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Table 10-4. Test acceptability requirements for 10-day NAF and 96-hr SBM tests with L.
plumulosus. Requirements listed are consistent with those specified in ASTM E 1367-92
and subsequent updates (E 1367-99)
Ten-day NAF and 96-hr SBM toxicity tests should usually be considered unacceptable if one or
more of the following occurred:*
• A 10-day NAF and 96-hr SBM toxicity tests are unacceptable if more than a total of 10%
of the control organisms die, or if the coefficient of variation (CV) of control survival is
equal to or greater than 40%.
• All test chambers were not identical.
• Test organisms were not randomly or impartially distributed to test chambers.
• Required reference standard was not included in the test.
• All test animals were not from the same population, were not all of the same species, or
were not of acceptable quality.
• Amphipods from a wild population were maintained in the laboratory for more than two
weeks, unless the effect of prolonged maintenance in the laboratory has been shown to
have no significant effect on sensitivity.
• The test organisms were not acclimated at the test temperature and salinity at least 48
hours before they were placed in the test chambers.
• Temperature and dissolved oxygen concentrations were not measured
*Note: These guidelines are not identical to those listed ASTM E 1367 in part because some acceptability guidelines
listed in ASTM E1367-92 are not applicable or practical for the NAF/SBM toxicity testing program.
10.7 Reference Tests
10.7.1 A single toxicity test will be used to determine satisfactory laboratory performance
and to determine whether an NAF or SBM can be discharged as it adheres to drill
cuttings. The reference toxicant for the NAF test will be either a Cie-Cig internal
olefin reference standard or a Ci2-Ci4 or Cg ester.
10.7.2 The reference toxicant for the SBM testing program will be a Cie-Cig internal olefin
SBM which has also been specified for determining pass/fail for SBMs. The Cie-Cig
internal olefin SBM is a 65/35 blend, proportioned by mass, of hexadecene and
octadecene, respectively (see 40 CFR 435.1 l(rr).
10.7.3 These reference toxicity tests will be conducted in conjunction with all NAF or SBM
tests to discern possible changes in the condition of the L. plumulosus population used
in testing. The reference toxicant test must be conducted concurrently with each
sample or batch of samples and at a minimum should be conducted at least monthly.
Control charts of this reference standard should be maintained to perform statistical
analyses, help understand the inherent variability in the reference test, and for long-
term quality control. Test conditions for the reference test should follow the
experimental conditions presented in Table 3.
10.7.4 The reference toxicant test should be performed concurrently-and under the same
conditions as the NAF or SBM test.
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10.7.5 The reference toxicant test should be conducted so that control limits (typically set at
2 standard deviations) can be established (U.S. EPA, 1994).
10.7.6 If the reference test LCso falls outside of this range of control limits generated on the
most recent test data points, then the sensitivity of L. plumulosus and the credibility of
the test results are considered suspect. In this case, the test procedure should be
examined and the test repeated with a different batch of amphipods. A sediment test
should not automatically be judged unacceptable if the reference test LCso falls
outside the expected range or if the control in the reference toxicity test exceeds 10%.
The width of the control limits and all performance criteria listed in Table 4 should be
considered when determining the acceptability of a given NAF or SBM test.
10.8 Interpretation of Result
10.8.1 Procedures presented in this test method are used to calculate point estimates, or LCso
values. The LCso value and 95% confidence limits of the NAF tests should be
calculated on the basis of milligrams of NAF per kg dry control sediment (mg/kg) and
amphipod mortality.
10.8.2 The LCso value and 95% confidence limits of the NAF tests should be calculated on
the basis of milliliters of NAF per kg dry control sediment (ml/kg) and amphipod
mortality.
10.8.3 A variety of methods can be used to calculate an LCso value and its 95% confidence
limits, including probit, moving average, trimmed Spearman-Karber and Litchfield-
Wilcoxon methods (ASTM E 1367-99). The method used should take into account
the number of partial kills, the number of test chambers per treatment (5), and the
number of amphipods per test chamber (20).
10.8.4 The only NAF-drill cuttings that can be discharge are cuttings coated with NAFs that
are as toxic or less toxic, but not more toxic, than the reference NAF (Cie-Cig internal
olefin or Ci2-Ci4 or Cg ester) as measured by LCso. This limitation is expressed in 40
CFR435.13, 435.15, 435.43, and 435.45.
10-day LCso Reference Material [10-3]
10-day LC50 NAF ~
10.8.5 The only SBM-drill cuttings that can be discharge are cuttings coated with SBMs that
are as toxic or less toxic, but not more toxic, than the reference SBM (Cie-Cig internal
olefin or Ci2-Ci4 or Cg ester) as measured by LCso. This limitation is expressed in 40
CFR435.13, 435.15, 435.43, and 435.45.
96-hr LCso Reference Drilling Fluid [10-41
10-day LC50 SBM ~ '
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10.8.6 The NAF or SBM data should be interpreted by comparing to the reference toxicant
test LCso value and whether it exceeds the limitations in 40 CFR 435.13, 435.15,
435.43, and 435.45.
10.9 References
10.9.1 American Society for Testing and Materials (ASTM). 1997. Standard Guide for
Conducting 10- day Static Sediment Toxicity Tests with Marine and Estuarine
Amphipods. E 1367-92. Annual Book of ASTM Standards, Vol. 11.05.
10.9.2 American Society for Testing and Materials (ASTM). 2000. Standard Guide for
Conducting 10- day Static Sediment Toxicity Tests with Marine and Estuarine
Amphipods. E 1367-99. Annual Book of ASTM Standards, Vol. 11.05.
10.9.3 Emery, V.L., D.W. Moore, B.R. Gray, B.M. Duke, A.B. Gibson, R.B. Wright and
J.D. Farrar. 1997. Development of a chronic sublethal sediment bioassay using the
estuarine amphipod Leptocheirusplumulosus (Shoemaker). Environ. Toxicol. Chem.
16:1912-1920.
10.9.4 McGee, B.L. DJ. Fisher, L.T. Yonkos, G.P. Ziegler and S. Turley. 1999. Assessment
of sediment contamination, acute toxicity, and population viability of the estuarine
amphipod Leptocheirus plumulosus in Baltimore Harbor, Maryland, USA. Environ.
Toxicol. Chem. 18:2151-2160.
10.9.5 Schlekat, C.E., B.L. McGee and E. Reinharz. 1992. Testing sediment toxicity in
Chesapeake Bay with the amphipod Leptocheirus plumulosus: an evaluation.
Environ. Toxicol. Chem. 11:225-236.
10.9.6 Suedel, B.C. and J.H. Rodgers, Jr. 1991. Variability of bottom sediment
characteristics of the continental United States. Water Res. Bull. 27:101-109.
10.9.7 U.S. EPA, 1994. Methods for Assessing the Toxicity of Sediment-associated
Contaminants with Estuarine and Marine Amphipods. EPA/600/R-94/025. USEPA
Office of Research and Development, June 1994.
10.9.8 U.S. EPA, 2001. Effluent Limitations Guidelines and New Source Performance
Standards for the Oil and Gas Extraction Point Source Category; OMB Approval
Under the Paperwork Reduction Act: Technical Amendment, Federal Register,
Volume 66, Page 6849-6919, January 22nd.
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