EPA 910-R-96-001
United States
Environmental Protection
Agency
Region 10
1200 Sixth Avenue
Seattle WA 98101
Alaska
Idaho
Oregon
Washington
Office of Environmental Assessment
April 1996
Microscopic Paniculate
Analysis (MPA) for
Filtration Plant Optimization
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Microscopic Participate Analysis (MPA)
For Filtration Plant Optimization
prepared by
Stephanie Harris1, Carrie Hancock1, & Jay Vasconcelos1
April 1996
1U.S. EPA Manchester Laboratory, 7411 Beach Drive East, Port Orchard, WA 98366
2CH Diagnostics & Consulting Service, 214 S.E. 19th Street, Loveland, CO 80537
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A cknowledgements
We wish to thank the many microbiologists and others who actively participated in the development and review of the
USEPA consensus method entitled "Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization".
Dr Susan Boutros
Environmental Associates
1185 East Main
Bradford, PA 16701
Rick Brinkman
250 North 5th St
Grand Junction, CO 81503
Carrie Howe-Carlson
Microsearch Laboratory
2783 Webster Drive
Grand Junction, CO 81503
Dr Jennifer Clancy (Randi McCuin)
Clancy Environmental Consultants
PO Bos 314
St. Albans,VT 05478
Jill Cunningham (Deborah Wayman, Deanna Crump)
Grants Pass Water Lab
558 ME "F"
Grants Pass, OR 07527
Rick Danielson (Craig Johnson)
BioVir Laboratories, Inc.
685 Stone Rd. # 6
Benicia, CA 94510
Michelle Eisenstein
Johnson State College
Chemical Hygiene Office
Johnson, VT 05656
Scott Tighe (Sharon Hallock, Brad Eldred)
Analytical Services, Inc.
PO box 515
Williston, VT 05495
John Rae
2855 Mesa Rd.
Colorado Springs, CO 80904
Dr. Frank Schaefer, III
USEPA/NERL/HERD/BARB
26 W. Martin Luther King Way
Cincinnati, OH 45268
We also wish to thank those who contributed materials and talent to design the cover page.
U.S. EPA Region 10 Graphics Department
BioVir Laboratories, Inc.
Eugene Water and Electric Board
Disclaimer
Mention of any trade names of commercial products does not constitute endorsement or recommendation for
use by the U.S. Environmental Protection Agency.
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Microscopic Participate Analysis (MPA) for Filtration Plant Optimization
Table of Contents
Introduction 1
Sample Collection '. 2
1.0 General Overview 2
2.0 Sample Equipment and Materials 2
3.0 Sample Collection Parameters 2
4.0 Sample Collection Procedure 3
5.0 Sample Volumes and Water Quality Parameters 4
Filter Processing and Analysis , 6
6.0 Equipment 6
7.0 Supplies 6
8.0 Processing Reagents 7
9.0 Paniculate Extraction 7
Palmer-Maloney Counting Cell 8
10.0 Subsample Examination 8
11.0 Microscopic Analysis 9
12.0 Whipple Grid Calibration 12
13.0 Centrifugate Pellet Measurement 12
14.0 Recording of Results & Procedural Parameters 13
15.0 Interpretation of Results 13
16.0 Analyst Qualifications 15
17.0 Standards of Identity 16
18.0 Quality Assurance 17
References for Internal Document 20
References for Microscopic Identification 20
Appendix 1 Microscope Alignment and Adjustment 22
Appendix 2 Use of Electronic Particle Counter 26
Appendix 3 Sample Data Forms and Report Forms 29
Appendix 4 Formulation ofMcFarland Standards 33
Appendix 5 Figures for Document 34
Appendix 6 Sample Calculation 39
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Microscopic Particulate A nalysis (MPA) for Filtration Plant Optimization
Introduction
With enactment of the 1986 Amendment to the
Safe Drinking Water Act (SDWA), the EPA has
promulgated new regulations for filtration and
disinfection of public water systems using surface water
or groundwater under the direct influence of surface
water (GWUDI). Those systems identified as surface
water or GWUDI must demonstrate a 3 log (99.9 %)
removal of Giardia and 4 log removal of virus particles
through a combination of filtration and disinfection.
Detection of Giardia cysts and Cryptosporidium
oocysts in surface water cannot be used to assess
treatment plant performance because of their low
concentrations, intermittent occurrence, and limitations
of the currently available technology. Because Giardia
and Cryptosporidium may occur at concentrations
sufficient to cause disease but not consistently
numerous enough to assess filtration performance,
surrogates for filtration efficiency have been and
continue to be developed.
Performance of water treatment plants can be
evaluated by a number of methods, including turbidity,
particle counts, and Microscopic Particulate Analysis
(MPA). Simultaneous use of more than one evaluation
technique may be appropriate, not only for research but
also for plant operation. Particle counting and turbidity
data must be used with caution because flocculated
particles could give false values compared with cyst
reduction and overall plant performance. Several
investigators have found that MPA can provide
information on the effectiveness of water treatment
processes for removing paniculate matter (1, 2).
Electronic particle counting, by itself, does not provide
the operator with critical information about the type and
number of organisms encountered.
MPA, including particle sizing, is performed
on drinking water systems where some form of
treatment, chemical or physical, exists between the
natural water source and its distribution to the public.
This analysis compares the type, size and quantities of
bioindicators and particles found in the raw water to
those found in the finished, or treated, water. This
method can be used to evaluate filtration efficiencies, as
log reduction, of conventional filtration systems, as
well as the on-site evaluation of alternate filtration
technologies.
This method can be used to identify certain
groups of microorganisms, 1 to 600 micrometer (/urn) in
size, which normally only occur in raw water as
opposed to finished waters and whose presence, in the
finished water, may indicate some breakthrough or
growth in the filter beds. These important
microorganisms, also called bioindicators, include
diatoms, algae, Giardia. coccidia, plant debris, pollen,
rotifers, crustaceans, ameba, nematodes and
insects/larvae. Comparison of the quantitative numbers
of these bioindicators in raw and finished water can
also assist in the over-all evaluation of filtration
efficiency and may provide information critical to the
optimization of the filtration plant beyond simple
turbidity reduction or particle counting by
instrumentation.
Historically, water treatment professionals
have relied on chemical and physical measurements to
assess water treatment plant performance; obviously
these assessments are not adequate because numerous
outbreaks of Giardia and Cryptosporidium have
occurred during periods where the plant met all
federally required performance criteria. An eclectic
approach using several tools, all of which measure
different aspects of plant performance, for the
assessment of filtration efficiency along with a
thorough understanding of the particular plant design
and operation helps avoid the inadequacy of simplistic
solutions for explaining the complex interactions of
water treatment.
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Sample Collection
1.0 General Overview: High Volume Filter (HVF) samples for surface water MPA are collected from the raw
water before it enters any phase of water treatment and from the finished water just prior to disinfection and
distribution. Evaluation of each filter bed or a composite of the effluent water is optional, and the chosen option
should be noted in the final report. However, blending of each filter bed into a composite many prevent
identification of individual filter bed inadequacies. The collection site should be selected to avoid stratification
of the pipes.
2.0 Sample Equipment and Materials
2.1 Sampling device consists of the following parts (refer to Figure 1 and 2)
2.1.1 Six foot inlet hose, preferable disposable, with backflow preventer (Watts No. 8)
2.1.2 Pressure regulator (Watts 26A), or equivalent, plus pressure gauge, 0-100 psi
2.1.3 Proportionating injector (for chlorinated water), Model 203 B.T. injector, 100-15P-87, or
equivalent. (Dema Engineering) (For chlorinated samples only).
2.1.4 Commercial Filter model LT-10 filter housing (9499-5015)
2.1.5 Water flow meter, readable in gallons or liters.
2.1.6 Flow control valve (limiting flow orifice) rated at 1.0 gallon per minute (gpm) for finished
water; 0.5 to 1.0 gpm for raw water. (Rationale for this modification is to allow collection
for a longer period prior to plugging of the filter in high turbidity waters)
2.1.7 Discharge hose
2.1.8 Pump, for non-pressurized sources
2.1.9 Miscellaneous brass, or PVC, fittings for unit assembly
2.1.10 Optional peto tube installed at sampling port is recommended to reduce problems caused by
flow dynamics in the pipe
2.2 Sampling Materials
2.2.1 Ten inch, 1 ^m nominal porosity, polypropylene, yam-wound, cartridge filter, Commercial
Honeycomb filter tube (M39R10A).
2.2.2 Whirl pak plastic bags (5.5" x 14") or zip loc heavy duty quality freezer bags
2.2.3 Sanitary gloves
3.0 Sample Collection Parameters
Note: Below are recommendations for typical treatment systems. Large, or atypical, systems where retention
occurs, may require alteration of these recommendations to provide accurate log reduction values. Moreover,
electronic particle count collection sites may vary from MPA collection points. If performing only electronic
particle counts, the sample might be more appropriately collected directly from the source, particularly if
presedimentation basins are an integral part of the treatment plant. The holding time associated with
presedimentation basins, may allow for settling of participates and may adversely influence log reduction
values.
3.1 Raw surface water should be sampled prior to chemical addition and after any presedimentation
basins (if no chemicals were added prior to presedimentation). The main objective in raw water
sampling is to collect a sample representative of the water entering the treatment system; therefore,
if recycling operations are practiced, the raw water should be sampled after the recycling input. Such
sampling should allow adequate time for mixing of recycling input prior to sampling. If collection
at the source is not possible, final report must "qualify" sample
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
3.2 Finished water should be sampled after the filtration system and prior to chlorine addition, if possible.
Sodium thiosulfate (final concentration 50 mg/1) is injected into samples that cannot be collected prior
to chlorination. Samples are collected prior to post treatment storage to provide a more accurate
evaluation of the filtration system. Evaluation of log reduction in large treatment plants with post
filtration holding tanks, may be difficult, given the propensity for algal growth in these circumstances.
3.3 Treatment plant evaluation. The raw water sampling should be initiated before the finished water
sampling. The amount of time elapsed between the beginning of raw sampling and the beginning of
finished sampling should be equivalent to the detention time of the system. To accurately assess
treatment efficiency, finished water sampling should encompass a full cycle run or for a 24 hour
sampling, including at least one backwash in the sampling.
3.4 Pressure over the filter face should be set at 10 psi, using the in-line pressure gauge and meter
4.0 Sample Collection Procedure (Flow chart, Figure 3)
4.1 Cleanliness- before each sample collection the hose and filter housing must be washed with hot water
containing a mild detergent and bleach solution; rinse with hot water followed by particle free water
(see 7.12). If this cannot be done, run a minimum of 50 gallons of sample water through the sampling
equipment prior inserting a new filter. Do not touch the filter with bare hands, use sanitary gloves or
the plastic cover the filter is wrapped in.
4.2 Connect sampling unit to pressure source and pump in the direction of flow indicated on filter housing.
Flush the unit without a filter for 3 - 5 minutes with the source water to be sampled.
4.2.1 Non-Pressurized Sources: Small 1-5 gallon per minute battery operated bilge pumps or
electric or gas powered centrifugal pumps may be used. Be sure to put the sample intake in
a location where the least amount of bottom sediment will enter into the sampling filter
giving a. distorted view of the sample. If possible, install the pump downstream (on the
effluent end) of the filter to eliminate the potential for cross-contamination of samples. Note:
Collect sample as near to intake site as possible. If intake is near the bottom and in fact
draws in bottom sediments, then collection here is appropriate.
4.3 Record the date, time of day and gallon reading from the water meter before and after sampling.
Document the name and location of each sample point, sampling site (raw or finished) and type of
treatment.
4.4 Insert filter in the housing and tighten housing. Make sure "0" ring is in place. Turn water on slowly
with the unit in an upright position. Invert unit to make sure all the air within the housing is expelled.
When the housing is full of water, return unit to upright position and increase flow up to 3.81pm (1
gpm). Measure flow rate by either timing the meter rate or by timing flow rate into a calibrated
bucket. Maintain 1 gpm (3.8 Lpm) throughout the sampling period. Exceptions to the 1 gpm rate are
given in 5.1. If 0.5 gpm flow control valve is being used follow the directions except that the flow
rate will be less and meter may not register so measuring the flow using a calibrated container may
be necessary.
4.5 Check reading on pressure gauge. Adjust pressure gauge to 10 psi, if needed..
4.6 Information on the sample volume and water quality parameters should be included on data sheet (See
section 5.0)
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
4.7 When sampling is complete, shut off flow, record stop time and final meter reading or time flow rate
into calibrated bucket and average with flow rate measured in 4.4. Subtract the initial reading from
the final reading and record the total volume collected.
4.8 Turn off the faucet or pump and disconnect the hose from incoming water source. Maintain the inlet
hose level above level of opening on outlet hose to prevent backwashing and loss of paniculate matter
from the filter. Pour residual water from filter holder into the ziploc (whirlpac) bag.
4.9 Remove the sampling cartridge with the plastic cover or sanitary latex gloves. Do not touch with bare
hands. Place filter in the heavy duty quality ziploc (whirlpac) bag and seal.
4.10 With permanent marker record the sample identification, gallons sampled, collection dates and times,
collector's name and water quality parameters directly on the bag or on a waterproof label. Place the
first bag containing the filter into the labeled ziploc bag. Make sure both bags are sealed to prevent
leakage.
4.11 If immediate shipping is not possible, the sample should be stored in a 1 - 5° C refrigerator, until
shipping, within limits set forth in 4.14.
4.12 Place freezer cold packs in the shipping container. Place insulating material between the filter and
cold packs to prevent HVF sample freezing. Samples that arrive at the laboratory frozen, should be
rejected, discarded and resampling requested. Place data sheet containing recorded information in a
sealed plastic bag and ship with the filters.
4.13 Ship by overnight delivery service to the analytical laboratory.
4.14 Samples must be processed within 96 hours of initiation of sampling.
5.0 Sample Volumes and Water Quality Parameters
5.1 Raw water: Sampling unit should be allowed to run for a 12 to 24 hour period in which time a
minimum volume of 100 liters (27 gallons) should be filtered. The ideal volume is the amount
equivalent to a complete day of production. If the filter becomes clogged or plugged due to highly
turbid waters, terminate sampling and record the volume collected to this point. If the raw water
source is known to have high turbidity, the sampling flow rate may be lowered to < 1 gpm to collect
a sample over a longer time period thus obtaining a sample more representative of the raw water
quality.
5.2 Finished water: Minimum 1000 liters or 264 gallons. Collection period should encompass a full
cycle run, or for 24 hours, including at least one backwash cycle. Backwash cycles can occur at the
initiation of the sampling period. If multiple filter beds are present in the filtration plant, a composite
sampling is recommended initially, although later evaluation of individual filter beds is an option.
Additionally, if requested, discrete sampling points at ripening, middle of cycle and after backwash
may be added to assist with plant operation.
5.3 Water Quality Parameters: Measurement of certain water quality parameters should be included
in the sample data form for both raw and finished water. Among these should be total and free
chlorine residual, temperature, pH, turbidity, and operational parameters of the WTP (pretreatment,
filtration, disinfection) and water source. Microbiological testing, such as total and fecal coliform and
heterotrophic plate count, are optional.
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
5.4 Chlorinated Samples: Try to sample water prior to any chlorination. If chlorinated water must be
sampled, an injector system will need to be installed to add a sodium thiosulfate solution to denature
the chlorine. Add sodium thiosulfate solution via the injector system to produce a final concentration
of 50 mg/L. Setting the injector system to produce a 1:100 dilution of 0.5% sodium thiosulfate stock
solution will result in a final concentration of 50 mg/L. Details on the operation and use of
proportioner pumps and injectors can be found in Standard Methods for the Examination of Water
and Wastewater. Section 95IOC, "Virus Concentration from Large Volumes by Adsorption to and
Elution from Microporous Filters (Proposed)," 18th ed., 1989, pp. 9-105 to 9-109. Model 203 B.T.
injector, 100-15P-87 special tip, Dema Engineering, or equivalent, may be used. Alternatively, a
peristaltic pump or electric pump can be used to inject the sodium thiosulfate.
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Microscopic Particulate A nalysis (MPA) for Filtration Plant Optimization
Filter Processing and Analysis (Fig 3,4)
6.0 Equipment
6.1 Large capacity centrifuge, refrigerated recommended
6.2 Large capacity swing-bucket rotor (90 °), 1-6 liter/run
6.3 250 mL conical bottom bottles with screw caps or 490 mL glass conical bottles
6.4 15 mL conical graduated centrifugate tubes
6.5 graduated cylinder
6.6 1 - 5° C refrigerator
6.7 50 mL conical graduated centrifuge tubes
6.8 Stomacher lab blender- model 3500 (optional)
6.9 Vortex tube mixer
6.10 Aspiration flask and vacuum source with 0-30 psi gauge
6.11 Pipet aid, syringe or bulb
6.12 Motorized multivolume microliter pipet (Rainin edp plus) or manual equivalent
6.13 Hollow glass tubes (ca 1/4" bore)
6.14 Brightfield, phase contrast, differential interference contrast (DIG) or Hoffman modulation optics
(HMO) capable microscope equipped with 10,40 and 100 X objectives. A 35 mm camera or video
camera attached, is optional
6.15 Manual or electronic differential counter (10 gang)
6.16 Non-drying immersion oil
6.17 Single place hand held counter
6.18 Palmer-Maloney counting chamber available from Wildlife Supply Company, catalog # 1803-B20,
specify glass model. (301 Cass St. Saginaw, MI, 48602)
6.19 Sedgewick-Rafter Counting Chamber (optional)
7.0 Supplies
7.1 Whirl pac bags, 5.5 x 15", sterile, or heavy duty ziploc bags. For filter transportation
7.2 Polypropylene yarn woven filter tubes (M39R10A, Commercial Filter, Lebanon, IN)
7.3 Sanitary gloves
7.4 Pan or tray, stainless steel or glass, autoclavable
7.5 4 liter beakers, autoclavable (glass or plastic)
7.6 Scalpel handles, autoclavable
7.7 Scalpel blades, sterile
7.8 Disposable glass pipets, sterile
7.9 Pasteur pipets, sterile
7.10 10 % buffered formaldehyde, pH 7.0
7.11 Polysorbate 80
7.12 Particle-free water (deionized, distilled or reverse osmosis water, passed through a 0.22 urn filter)
should contain less than 100 particles/ml (2 ^m or larger)
7.13 Clear fingernail polish
7.14 3.5 L capacity Stomacher bags (Seward medical, Tekmar Co)
7.15 2 L beakers
7.16 Non-drying immersion oil
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
8.0 Processing Reagents
8.1 Wash water (non-sterile)
8.1.1 Sterile Erlenmeyer flask (1,2 or 4 L)
8.1.2 Particle free water (7.12)
8.1.3 Sodium citrate (optional, if iron present)
8.1.4 0.01 % polysorbate 20, add and mix immediately prior to use (optional)
8.1.5 Mix these in the following proportions:
Wash-Water Proportions
LL 2L 4L
Sodium citrate (Optional) 5.0 g lO.Og 20.0g
0.01 % polysorbate (Optional) 10.0 ml 20.0 ml 40.0 ml
Particle-free water (Quantity Sufficient to make) l.OL 2.0 L 4.0 L
Final pH to 5.5 - 7.5
8.1.6 Use of 0.85 % NaCl or other buffering agent is optional
8.1.7 Record constituents of wash water on laboratory bench sheet
9.0 Participate Extraction: The filter cartridge is handled aseptically. All glassware and other equipment is
mechanically scrubbed, rinsed in particle-free water and autoclaved or chemically sanitized. Sanitary gloves
are worn during processing.
9.1 Remove filter from the ziploc/whirl pac bag and place in pan.
9.2 Record the color of the filter and any other notable physical characteristics.
9.3 Rinse the bags with particle-free water. The rinse water is retained in a beaker.
9.4 The filter fibers are cut length-wise to the filter core and separated into a minimum of 6 equal portions.
Each portion is washed sequentially in 3 consecutive 1.0 liter volumes of wash water. Begin with the
cleanest fibers and proceed to the dirtiest. Washing consists of vigorous kneading and swirling
motion. Minimum total wash time of fibers should be 30 minutes. The fibers are wrung out into a
collection beaker by placing them in individual interlocking bags which have one comer snipped off
to allow for drainage.
9.5 Alternatively, a Stomacher lab blender (model 3500) may be substituted for handwashing. The filter
is cut length-wise to the core and after loosening the fibers, place all of the fibers into a single
stomacher bag. To insure against bag breakage and sample loss, place the filter fibers in the first
stomacher bag into a second stomacher bag. Add 1.75 L of wash water to the fibers. Homogenize
for 2- five minute intervals. Between each homogenization period, hand knead the filter material to
redistribute the fibers in the bag. Wring the fibers out to express as much of the liquid as possible and
place them in another stomacher bag containing 1.25 liters of wash water. Repeat for 2- five minute
homogenization periods. Wring the fibers to express as much of the liquid as possible before
discarding.
9.6 The 2 aliquots of wash water, bag rinse water and residual filter water obtained from the filter housing
are combined in one 4 liter beaker.
9.7 Record the volume of the total paniculate solution.
Note: The use of Immunofluorescent Assay for Giardia & Cryptosporidium is optional at this time, but is required
if Giardia & Cryptosporidium reporting is included in the analysis. If IFA is chosen, follow the latest method
publicized in the federal register. The fibers from those samples for which IFA is being done in addition to
MPA will be washed a second time in the IFA prescribed eluting solution (Federal Register 1994). An FA
eluting solution was developed because more Giardia cysts and Cryptosporidium oocysts could be recovered
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
during the washing process than by using particle-free water alone. However, many non-encysted organisms
subjected to the FA eluent deteriorate and cannot be recognized or identified for MPA. Therefore, a paniculate
extraction scheme will need to be followed so that both MPA and FA can be done from the same sample. The
particle-free water/particulate solution from the initial wash is halved for MPA and FA. The half retained for
IFA is combined with half of the secondary wash solution to provide a known quantity of IFA/particulate
solution.
Using Palmer-Maloney Cell
The Sedgwick Rafter (S-R) may be used in addition to the PMCC for enumeration of larger zooplankton if long
working distance 20x or 40x objectives are available and if written records of calculation and methodology following
Standard Methods are kept.
10.0 Subsample Examination: The total particulate solution represents a solution of the participates recovered
from the number of gallons sampled; therefore, an accurate record of the number of gallons sampled is a
prerequisite for the calculations needed to do any further processing. Convert the number of gallons to liters.
10.1 Thoroughly mix the total particulate solution by pouring it back and forth in glass containers of
sufficient size to hold the whole sample, or alternatively, use a sterile stir bar and magnetic plate and
mix for 10 minutes. After mixing, immediately remove a 200 ml aliquot and place into a 250 ml
conical centrifuge bottle. Vortex for IS seconds.
10.2 Immediately after mixing, withdraw a 0.1 mL subsample with an calibrated Eppendorf, or equivalent,
pipette.
10.3 Inject the subsample into the Palmer-Maloney counting chamber by introducing the sample with the
pipette into one of the two 5-mm channels on the sides of the chamber with the cover slip in place.
10.3.1 Calculate and record the liter equivalent of the withdrawn subsample using this proportion
ratio:
Total Volume in 250 ml conical centrifuge bottle = 0.1 mL
Liter equivalent in centrifuge bottle X
where: X = liters equivalent of 0.1 ml subsample
Liter equivalent in centrifuge bottle is calculated from this proportion ratio:
Total volume of particulate solution (in ml) = 200 ml aliquot
Total # of Liters sampled Liter equivalent in centrifuge bottle
Note: If returning to this step for a second time, skip the above calculation; double the previously calculated
Liter equivalent in centrifuge bottle (triple if a third 200 ml aliquot is added), then subtract the liter equivalent
of previously withdrawn 0.1 ml subsample (10.7.1).
Note: If previous history of this treatment plant warrants it, centrifugation of the entire wash water may be
performed.
10.4 View the subsample at lOOx magnification. If >10 plankters (plural for individual plankton
organisms) are present per field of view, proceed to section 11.0.
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
10.5 If <10 plankters per field of view are present, centrifuge the bottle at 1050 x g for 10 minutes.
Alternatively, if the turbidity of the paniculate solution examined is comparable to a 1.0 McFarland
standard, this will provide an acceptable density of particulates in most instances. (See McFarland
Standards in Appendix 4.)
10.6 Aspirate the supernatant down to 4cm above the bottom sediment or the bottom of the bottle, this
should provide approximately 50 ml. Measure and record the volume of combined cenlrifugate and
supernatant.
10.7 Thoroughly mix the solution by vortexing for 15 seconds and immediately withdraw a 0.1 ml
subsample for charging the Palmer -Maloney counting chamber.
10.7.1 Calculate and record the liter equivalency of the 0.1 mL aliquot using this proportion ratio:
(For example of calculations see Appendix 6.)
Volume of Remaining Particulate Solution (in mL) = 0.1 mL
Remaining Liter Equivalency X
where:
X = liter equivalent of 0.1 ml subsample
Vol. of Remaining Particulate Solution = Vol.
recorded in 10.6.
Remaining Liter Equivalency = Liter equivalent in centrifuge bottle calculated in
10.3.1 less the liter equivalency of previously withdrawn 0.1 mL subsample(s).
10.8 If >10 plankters are present per lOOx magnification field of view, proceed to section 11.0.
10.9 If <10 plankters per field of view are present, add an additional 200 ml aliquot of the total paniculate
solution by repeating steps 10.1 to 10.9 until the plankter density is correct. Alternatively, if the
turbidity of the paniculate solution examined is comparable to a 1.0 McFarland standard, this will
provide an acceptable density of particulates in most instances. (See McFarland Standards in
Appendix 4)
NOTE: Frequently, raw surface water may be examined without centrifugation after filter washing;
whereas, finished water often requires several centrifugation steps. Sometimes samples are over-
concentrated (too dense for microscopic visualization due to overlapping) and need to be diluted. If
further dilution is necessary, remember to include this in the calculations in 10.3.1 or 10.7.1.
Occasionally, interfering amorphous debris (ex: detritus or flocculent from a conventional treatment
plant) prohibits a density of 10 plankters per field and these samples must be examined as described
in section 11.0 at a particle density as dense as possible but without overlapping particulates that could
obscure visualization of the plankters.
11.0 Microscopic Analysis
11.1 After application of sample, allow a 10 minute settling period before counting. The entire Palmer-
Maloney counting chamber is systematically examined at a minimum of lOOx magnification using
phase optics, brightfield, DIC or HMO. Begin scanning the chamber at one edge and use an up-and-
down or a side-to-side scanning pattern (see Figure 5). If the distribution of organisms is random and
the population fits a Poisson distribution, the counting error may be estimated. If 100 units are
counted, the 95 % confidence limits approximate ± 20 %. (3,4).
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
11.1.1 Separate counts are made for each of the following size ranges: < 10 um, 10 -25 um, 25 -
100 um, 100 - 200 um, > 200 um and total number and should be recorded on the data sheet.
An ocular micrometer calibrated against a stage micrometer should be used when performing
particle sizing. All categories from 11.1.4, as well as amorphous debris and unclassified
biological material will be included in this count. Using a calibrated whipple grid (Appendix
1), count all particles, at a minimum of 100X magnification, present in a total of 20 - 30
whipple grid fields chosen randomly from the PMCC. Averaging methods are acceptable
with densities too high for accurate counts. Record the number of particles found in each
size range in each of the grid fields. Formula for calculation of # per 100 1 is in section
11.2.3 and 11.2.5.
11.1.2 Any microbiota seen that are not too numerous to count at a minimum magnification of lOOx
are identified following the Standards of Identity section.
11.1.3 Identification to the lowest level of taxonomic resolution known by the analyst is recorded.
11.1.4 Separate counts are made for each of the following categories: nondiatomaceous algae,
diatoms, plant debris, rotifers, nematodes, pollen, ameba, ciliates, colorless flagellates,
crustaceans, other arthropods and "other" (see Standards of Identity section). Counts are
made by the natural unit (clump count enumeration) method defined as follows: any
unicellular organism or natural colony is counted as one organism. Alternatively, total count
method can be used, defined as follows: each cell is counted as 1 organism. Making a total
cell count is time-consuming and tedious, especially when colonies consist of thousands of
individual cells. The natural unit is the most easily used system, however it is not necessarily
the most accurate because sample handling, collection or water treatment may result in
breakdown of organisms leading to inaccurate removal rates between raw and finished water
samples. Analyst must report the enumeration method used. Counts are made for those
categories of organisms that are not too numerous to count.
11.1.5 The counts for each category are extrapolated from the 0.1 mL aliquot used in the Palmer
Cell to numbers per 100 Liters as follows:
11.1.5.1 Calculate X using this proportion ratio: (For calculation example see Appendix 6.)
# of organisms = X
O.lmL V
where: # of organisms = the count from one category
V = volume from which 0.1 ml aliquot was withdrawn
(derived from 10.3.1 or 10.7)
11.1.5.2 Calculate the number of organisms per 100 liters for each category using this
proportion ratio:
X = number of organisms
liters equiv. of soln from 100 liters
which 0.1 ml aliquot was withdrawn
where:
X = calculation from 11.1.5.1
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Microscopic Paniculate A nalysis (MPA) for Filtration Plant Optimization
11.2 The common organisms, or particles that are too numerous to count at lOOx are counted in calibrated
Whipple grid fields at 400x. Specific calibration of each microscope used is essential (see 12.0).
11.2.1 As many Whipple grid fields as necessary to obtain a minimum of a 100 organism count are
observed. (95 % CL of approximately ± 20 %.) Use a single place hand held counter to
count the number of fields observed. Use a manual differential counter (10 gang) or 10 place
electronic tabulator to count the number of organisms observed in each of the categories
listed in 11.1.4. Count only those categories that were not counted in section 11.1.4.
11.2.2 Record the number of organisms or particles found in each category as well as the identity
of the organisms observed at the lowest level of taxonomic resolution known by the analyst.
11.2.3 Calculate the number of organisms or particles per mL in each category using the following
formula:
No./mL = C x 1000mm
A x D x F
where:
C = number of organisms counted for each category from 11.2.2
A = area of a field (Whipple grid image), mm2 (width2 - see 12.0)
D = depth of a field (P-M chamber depth = 0.4mm)
F = number of fields counted
11.2.4 Calculate the number of organisms in the remaining paniculate solution from which 0.1 mL
of solution with > 10 plankters per field was withdrawn as follows:
N_ = X
ImL mLs of remaining paniculate solution
where:
N = number of organisms per mL (calculated in section 11.2.3)
X = number of organisms in the remaining paniculate solution
11.2.5 Calculate the number of organisms per 100 liters using this proportion ratio:
X = No. of organisms
L 100 liters
where:
X = the number of organisms in the remaining paniculate solution calculated
in section 11.2.4.
L = liters equivalency of the remaining paniculate solution (Total number of
liters sampled less the liter equivalence of withdrawn subsamples).
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
12.0 Whipple Grid and Ocular Micrometer Calibration: Place a Whipple grid in an eyepiece of the microscope
and a stage micrometer that has a standardized, accurately ruled scale on a glass slide. The whipple disk has
an accurately ruled grid subdivided into 100 squares. One square near the center is subdivided further into 25
smaller squares. The outer dimensions of the grid are such that with a lOx objective and a lOx ocular, it
delimits an area of approximately 1 mm2 on the microscope stage. At 40x and with the ocular and stage
micrometers parallel and in part superimposed, match the line at the left edge of the Whipple grid with the zero
mark on the state micrometer scale. Determine the width of the Whipple grid image to the nearest 0.001 mm
from the stage micrometer scale. (APHA 1995)
13.0 Centrifugate Pellet measurement (Figure 4) Centrifugate pellet measurement may provide information about
the overall plant performance. However, it is not intended to be used as a sole method for determining filtration
efficiency.
13.1 If the liter equivalent in the 250 ml centrifuge bottle (10.3.1) is > 100 liters proceed to 13.1.1. If not,
proceed to 13.2.1.
13.1.1. Centrifuge the 250 ml bottle at 1050 x g for 10 minutes. Measure the Centrifugate pellet
volume. If volumes are below lowest graduation, mark a "dummy" set of tubes using water
injected from calibrated pipettes and compare to sample.
13.1.1.1. Calculate the volume of Centrifugate pellet per 100 liters using the
following proportion ratio:
Total Centrifufiate Pellet X
Liter equivalent in 250 ml centrifuge bottle = 100 liters
where:
Total Centrifugate pellet = volume of Centrifugate from 13.1.1
X = The volume of Centrifugate pellet per 100 liters
13.2.1 Remove a subsample from the remaining paniculate solution equivalent to 100 liters. For
example: if the total paniculate solution is 4,000 mL representing 400 liters sampled, 1,000
mL would be removed.
13.2.1.1 Pour the subsample into 50 mL tubes. Centrifuge at 1050 x g for 10 minutes.
Aspirate the supernatant down to 5 mL above the bottom sediment. The remaining
subsample may be poured on top of the Centrifugate pellet and remaining
supernatant in the 50 mL tubes. Centrifuge again at 1050 x g for 10 minutes. This
may be repeated until all of the 100 liter equivalent subsample has been centrifuged.
13.2.1.2 Aspirate the supernatant down to 5 mL above the Centrifugate pellet. Combine the
pellets and the remaining 5 mL of supernatant into one 50 mL tube. Centrifuge at
1050 x g for 10 minutes.
13.2.1.3 Record the Centrifugate pellet volume.
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
14.0 Recording of Results and Procedural Parameters
14.1 Field data should include the following:
14.1.1 Total water volume filtered in gallons
14.1.2 Water source identified as to type and location
14.1.3 Record the type of filtration, any pretreatment and kind of disinfection
14.1.4 Record both address and exact location of water source being evaluated
14.1.5 Date and time of sample device installation and removal
14.1.6 Name, address and phone numbers) of sampler(s)
14.1.7 Field measurements, such as turbidity, pH, conductivity, chlorine residual
14.1.8 Record use of sodium thiosulfate if applicable
14.2 Laboratory data should include the following:
14.2.1 Total volume of packed pellet .(centrifugate volume)
14.2.2 Number of each bioindicator from 0.1 ml sample aliquot. Proper significant figures should
be used in calculations. Refer to Standard Methods for Water and Wastewater. 19th ed for
details.
14.2.3 Number of particulates from each slide for the size ranges described in 11.1.1
14.2.4 Type of microscopy employed
- brightfield
- phase contrast
- other
14.2.5 Magnification of objective(s) used
14.2.6 Number of whipple fields per PMCC at 400 X or other magnification
14.2.7 Wash Solution constituents (e.g. Tween 20, Sodium citrate)
14.2.8 Sedgewick Rafter or Palmer Maloney Counting Cell used
14.2.9 Kind of count used; total or natural unit
15.0 Interpretation of Results: MPA for filtration plant optimization identifies and enumerates a subsample of the
organisms/particles eluted from a HVF waterborne paniculate sample collected through a 1 um nominal
porosity filter cartridge. Other paniculate measurements and observations are also reported. If raw and finished
samples are analyzed, an estimate of filtration plant efficiency can be determined. In addition, since biological
organisms are identified, potential filtration plant problems may be identified and can lead to optimization of
plant operation.
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
15.1 Filter Color. The one micrometer filter cartridge changes color during sampling depending on the
water's paniculate composition and color as well as the amount of water sampled. The cartridge color
can provide useful information about the general quality of water and can be used to make some
process control decisions. For example: an efficient water treatment plant will often have a brown raw
water sampling cartridge and a white finished water sampling cartridge. The presence of a green tinge
on only the finished filter cartridge may indicate the presence of algae growth within the filter beds.
15.2 Centrifugate pellet volume. The centrifugate pellet volume in ml per 100 liters is a direct
measurement of the final pellet of paniculate matter recovered from the sampling cartridge after
paniculate elution and centrifugation. The percent reduction or log removal between raw and finished
centrifugate pellet volume can be useful in interpretation of overall filtration plant efficiency.
However, it is important to realize that the volume of pellet can be strongly influenced by sampling
technique and other factors and therefore should not be used as the sole factor in determining filtration
efficiency. Treatment problems may be identified when finished sediment is greater in volume than
the raw sediment. Situations such as this may occur when excess treatment chemicals are used.
Percent reduction of centrifugate volume through the treatment system is calculated as follows:
15.2.1 % reduction = (raw centrifugate - finished centrifugate) x 100
Raw centrifugate
Log removal = Log (Raw Centrifugate) - Log (Fin. Centrifugate)
Example:
Raw Water = 3.0 mL per 100 liters
Finished Water = 0 J mL per 100 liters
% Reduction = 3.0 mL (in) - 0J mL (out) x 100 = 90%
3.0mL (in)
The above example would equal 1.0 log removal.
15.3 Amorphous Debris. This category lists the types of inorganic matter and non-living organic matter
(detritus) in order of predominance. Also included is the size range of these particulates observed
under the microscope. Particulates are reported in numbers per 100 liters, using proper significant
numbers (scientific notation is optional) as outlined in Standard Methods for the Examination of Water
and Wastewater (19 ed, 1-17)
15.4 Nondiatomaceous Algae through Other (refer to data sheet). These categories signify the different
types of organisms found and their respective numbers. Organisms are reported in numbers per 100
liters, using proper significant numbers. A qualitative approach can be taken for each category of
organisms or a more objective approach can be taken by examining the total organisms percent
reduction or log removal by the treatment system.
15.5 The percent reduction and log removal of organisms is calculated as follows:
15.5.1 Calculate the total number of organisms found in the raw water and in the finished water.
15.5.2 % Reduction = (Raw total - Finished total) x 100
Raw total
Log Removal = Log,0 (Raw total) - Log10 (Finished total)
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
15.5.3 Example:
Raw Water Total Organisms = 40,000,000/100 liters
Finished Water Total Organisms = 400,000/100 liters
% Reduction = 99 %, or 2 log removal. 99.9 % reduction is equivalent to 3 log
removal.
15.5.4 Percent reduction and log removal can be approximated for plants with more than one raw
water source if MOD or percent use records are kept for each source. Separate counts from
each source are weighted by the appropriate percentage to calculate a total influent count.
For example: A water utility filters 21 MOD; it is composed of 14 MOD (67 % of
total) of reservoir water and 7 MOD (33 % of total) from a river. Analysis of both
influent sources provides the following information on algal content:
1) Reservoir contains 50,000 per 100 liter
2) River contains 100,000 per 100 liter
The total influent count to be used in percent log removal (river and reservoir
combined) would be calculated as follows:
(50,000 x 0.67) + (100,000 x 0.33) = 60,000 algae per 100 liter.
15.5.5 . Occasionally organism numbers may increase in the finished water, and it is suspected that
either reproduction is occurring in the filter beds or another source of raw water is being
introduced that has not been accounted for such as backwash return water or some other in-
plant recycle operation. In such instances, information about the kind of organisms present
in the finished water may assist the plant operator in improving plant operation. Likewise,
the total centrifugate volume may increase in the finished water, or may demonstrate less log
removal than the organism removal. For example, flocculent may be passing through the
system. Identification of this substance microscopically, may provide useful information to
the plant operator.
16.0 Analyst Qualifications
Interpretation of results derived from the consensus method will depend upon numerous factors, the most
important of which will be the level of training and experience of the analyst(s) employing this technique.
16.1 Analyst should have a strong background in limnology and freshwater biology as well as an academic
background and/or training in parasitology, protozoology, phycology, invertebrate zoology and
bacteriology.
16.2 Analyst should have extensive experience with a light microscope with skills in brightfield, phase
contrast and DIC or HMC microscopy.
16.3 Analyst should have experience in examining a sufficiently large number of Surface Water MPA
samples.
16.4 A working knowledge of conventional treatment plants, slow and rapid sand filters, and alternative
filtration methods is essential to providing adequate interpretation of the results and recommendations
for controlling treatment plant conditions.
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
17.0 Standards of Identity
Diatoms:
Other Algae
Rotifers:
Plant debris:
Nematodes:
Pollen:
Diatoms are a group of algae which are distinctive because their cell wall is composed of
silica. This contributes to their ability to resist environmental, mechanical and chemical
insults. There are numerous species found in surface waters. They contain chlorophyll and
need sunlight to live and reproduce. The size of these organisms is dependent on the
nutritional quality of the water. Some species are known to be nuisance organisms because
they clog filtration systems. A preponderance of 1 or 2 species in the finished water
indicates possible reproduction or retention in the filter beds rather than the actual passage
through the filtration plant. It is important to categorize the diatoms presence as living
(containing internal structures) or dead (empty silica skeletal remains). The number of empty
or dead diatoms may be of interest in certain types of filtration plants (e.g. effluent from DE
filters frequently contains large numbers of empty diatoms).
This category is comprised of a large number of chlorophyll containing filamentous, colonial
and unicellular divisions of algae. Like diatoms, these genera of chlorophyll-bearing algae
require sunlight for their metabolism. Surface water contains more than 10,000 known
species with about 100 different species being commonly found. Diversity, abundance and
organism size are dependent on available nutrients, water temperature, time of year, and
other environmental and biological factors. Some species are known to be nuisance
organisms causing taste and odor problems. Some cause filters to clog and add color to the
water. Some species will reproduce in the filtration system and be present in the effluent.
This can usually be detected because there is an overall decrease in the variety of species and
an increase of only a few species between the raw and finished water.
A major taxonomic group with over 2500 species, of which more than 2375 species are of
fresh water origin. They are associated with a variety of habitats including small puddles,
damp soils and vegetable debris. They are also found associated with mosses, which can
often be found in or around ground water sources. The vast majority of rotifers encountered
are females ranging in size from 70-500 urn. Rotifer growth in filtration beds has been
suggested.
This group may be defined as either unidentifiable plant material containing chlorophyll or
undigested fecal detritus from herbivorous animals, usually muskrat and beaver. Plant debris
is very light weight material which is large in size (50 - 100 um). If the plant material is
fecal detritus, it can suggest that animals are present in the watershed and may shed cysts or
oocysts.
These include some 2,000 known free-living species found in fresh water. Some species
show an amazing ability to survive and thrive in aquatic habitats under a wide range of
ecological conditions. Benthic sediments of lakes and rivers can contain high numbers of
nematodes, as can sewage effluent. The top layer of soil can contain over 1 million
nematodes per square meter. Soil runoff is a major source of nematodes in source waters for
treatment plants. Nematodes and/or their eggs are common in healthy water sources.
Nematodes found in finished potable water do not portray a quality product to the public and
may also compromise the microbiological integrity of the drinking water. These organisms
seem to grow or reproduce in filter beds and distribution systems, so proper backwashing and
super chlorination of the filter beds, as well as, proper maintenance of the distribution system
should be conducted routinely.
This includes all microspores produced by plants. In the spring and fall, pollen is
everywhere, both airborne and waterborne. Because pollen can become trapped in the filter
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
cartridge during insertion of the filter or in the laboratory while the filter is being processed
for examination, it is only rarely useful for assessing filtration efficiency, by itself.
Ameba: These include the ameboid, flagellated and cyst stages ranging in size from 10 to 600 um.
This group is characterized by the formation of pseudopodia of one type or another. The
external surfaces of these ameba are usually very thin compared to the cell coverings of
ciliates and most flagellates. Most species are free-living and feed on bacteria, algae, other
protozoa and debris. Ameba are common in surface waters and proper filtration removes
them, but reproduction may occur in filter beds.
Ciliates: These free-living protozoa are very common. Ciliates are distinguished from other protozoa
by the presence of a macronucleus. Like amoeba, they feed on bacteria, algae, small
metazoa, other protozoa and debris. Proper filtration removes ciliates, but a few species may
reproduce in filter beds.
Colorless flagellates:
Although many flagellates are phototrophic, there are many colorless species that grow in
the absence of light if sufficient dissolved nutrients are available. They are common in
surface water and can be removed by filtration; however, some species may reproduce in
filter beds. Flagellates possessing chlorophyll are included in the algae category.
Crustaceans: These include all aquatic arthropods which have two pairs of antennae and are fundamentally
biramous. The vast majority of known species (>35,000) are marine, but approximately 1,200
are found in freshwater. Adults range in size from 250 to 500 um, with eggs from 50 to 150
um. Several species occur in healthy surface and ground water. Daphnia and Bosmina
species have been known to reproduce in very high numbers under the right environmental
conditions and cause filter clogging problems for water treatment plants. Finished and
distribution waters can contain large numbers of crustaceans. It is suspected that eggs will
hatch in the filter beds or pass through the filters and hatch in the distribution system.
Identification of crustaceans is often difficult because of fragmentation and observation of
only small portions of the organism.
Other Arthropods:
There are a large number of organisms, all with jointed appendages, in the phylum
Arthropoda. This category includes only the arthropods that are not classified as crustaceans
or those which are identifiable only to the phylum level due to the decomposed and
fragmented condition of the organism. Chironomid (insect) larvae and eggs are commonly
reported in surface waters as are arthropod pieces. Seen less frequently are other insects,
water mites and seed ticks.
Other: This category includes any organism seen that does not fit into the above categories.
Examples include, iron bacteria, fungal spores, gastrotichs and/or tardigardes.
18.0 Quality Assurance. Listed below is the minimum recommended QC to be followed to under a laboratory
QA/QC program. Documentation of testing is extremely important and careful records need to be maintained
at all stages of analysis. Additionally, users of this method should develop their own internal QA/QC, and
attempt to determine precision and bias at least at the analyst level for paniculate counts.
18.1. QC on equipment and supplies.
18.1.1 Large capacity high/low speed centrifuge (preferably refrigerated).
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
18.1.1.1 Equipped with swing bucket rotors. Records maintained on rotor(s) usage
at designated RPM. Post Manufacturer recommendation with regard to
life time hours on rotor.
18.1.1.2 Rotor speed checked with tachometer on a yearly basis.
18.1.1.3 Determine and record RPM necessary for each rotor to attain desired g
force. Post near centrifuge.
18.1.1.4 Annual PM agreement in force or internal maintenance protocol/records
in place.
18.1.2 Brightfield/phase-contrast/DIC/HMO microscope (Appendix 1)
18.1.2.1 Phase rings checked for each objective before each use period. Kohler
illumination adjusted for each objective for DIC/HMO.
18.1.2.2 Ocular micrometer (reticle) in place and calibrated against a stage
micrometer for each objective in use. Calibration data posted near
microscope. Re-check on an annual basis.
18.1.2.3 Microscope must be cleaned and optics realigned and adjusted on a
frequent schedule.
18.1.2.4 Annual PM agreement in force or internal maintenance protocol/records
in place.
18.1.2.5 Whipple grid used must be designed for installation into the laboratory's
microscope ocular and must be calibrated against a stage micrometer.
18.1.3 Stomacher brand (model 3500) laboratory blender.
18.1.3.1 Operated according to manufacturers recommendations. The use of the
blender is carefully timed to insure consistent washing of filter fibers.
Stomacher is properly adjusted, with set screws, to accept entire filter.
18.1.3.2 Stomacher unit is maintained and internal paddle cover is
cleaned/disinfected after each use with dilute detergent/bleach solution.
18.1.4 MPA sampling apparatus
18.1.4.1 Apparatus is cleaned with dilute detergent and bleach solution, rinsed
thoroughly with hot tap water, followed by a particle-free water rinse in
the lab. Apparatus is flushed with water in the field prior to inserting the
filter.
18.1.4.2 Water meter is periodically checked for accuracy by timing the rate of
flow into a measured gallon container.
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
18.1.5 Sample processing
18.1.5.1 All laboratory supplies used during sample processing are autoclaved or
chemically sanitized.
18.1.5.2 Particle-free water has been tested and shown to contain less than 100
particles per 100 ml.
18.2 Analytical QC
18.2.1 Analyst has available identification keys and pictorial atlases to assist in classification of
microbiota. (see reference list).
18.2.2 Strict adherence to the Consensus Method and the definition of Standards of Identity will aid
in maintaining intralaboratory and interlaboratory consistency.
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Microscopic Paniculate A Italy sis (MPA) for Filtration Plant Optimization
References for Internal Document
1. Vasconcelos, GJ. and S.I. Harris. EPA in-house document. Microscopic Particulate Analysis and
Particle Size Analysis for Determination of Filtration Efficiencies. EPA Manchester Environmental
Laboratory.
2. Hancock, C.M., J.V. Ward, K.W. Hancock, P.T. Klonicki and G.D. Sturbaum. Assessing Water
Treatment Plant Performance Using Microscopic Analysis (MPA). Proc AWWA WQTC Nov.
1994, San Francisco.
3. Palmer, C.M. and T.E. Maloney. 1954. A New Counting Slide for Nannoplankton. Amer. Soc. of
Limn, and Oceano. special pub. no. 21, Environ. Health Center, Public Health Service, Cincinnati,
Ohio.
4. Rashash, D. and D.L. Gallagher. 1995. An Evaluation of Algal Enumeration. Journal AWWA,
April 1995. 127-132.
References for Microscopic Identification
American Public Health Association, American Water Works Association, Water Environment Federation. 1995.
Standard Methods for the Examination of Water and Wastewater. Section 10200 F. Phytoplankton Counting
Techniques & color plates. 19th Edition, A.P.H.A., Washington, D.C.
AWWA (1995) Selected Problem Organisms in Water Treatment. M7 Operator's Identification Guide.
A;P.H.A., A.W.W.A., W.E.F. 1994. Section 9711 B. Immunofluorescence Method for Giardia and Cryptosporidium
spp. Standard Methods for the Examination of Water and Wastewater (Proposed). 19th Edition, A.P.H.A.,
Washington, DC, pp. 9-111.
Belehery, Hilary, 1979. An Illustrated Guide to Phvtoplankton. H.M. Stationery Office, London.
Boutros, S.N. 1993. Microscopic Particulate Analysis (MPA) in Studies of Ground water. Proc. AWWA WQTC,
Nov. 15-19,1992, Toronto, Ontario.
Brady, Nyle C. 1974. The Nature and Property of Soils. MacMillan Publishing, New York, 637 pp.
Federal Register, Monitoring Requirements for Public Drinking Water Supplies; Proposed Rule, February 10,1994.
40 CFR Part 141. Vol. 59, No. 28 Proposed Rules.
Foged, Niels, 1981. Diatoms in Alaska. Bibliotecha phvcoligica. J.Cramer Inder A.R. Gantner Verlag
Kommaniditgesellschaft.
Garnett, WJ., 1965. Freshwater Microscopy in U.S. Dover Publishing, Inc., N.Y., N.Y.
Lee, JJ., Hunter, S.H. and E.C. Bovee, editors. 1985. An Illustrated Guide to the Protozoa. Society of Protozoologists,
Lawrence, KS.
Lund, JWG and H. Canter-Lund, 1995. Freshwater Algae; Their Microscopic World Explored. Biopress Ltd.,
Bristol, England.
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Palmer, C. Mervin, 1962. Algae in Water Supplies. U.S. Dept. of Health, Education and Welfare, Public Health
Publication No. 657, Washington, D.C.
Pennak, R.W. 1989. Freshwater Invertebrates of the United States Protozoa to Mollusca. 3rd edition. John Wiley
and Sons, Inc., New York.
Pentecost, A. 1984. Introduction to Freshwater Algae. Richmond Publishing Co. Ltd., Richmond, England.
Pesez, Gaston, 1977. Atlas de Microscopic des Eaux Douces. 19 rue Augereau, Paris.
Prescott, G.W., 1954. How to Know the Freshwater Algae. In: Pictured Key Nature series, University of Montana
Press, Montana.
Smith, G.M., 1950. 2nd ed. Freshwater Algae of the United States. McGraw Hill
Book Co., Inc., N.Y.
United States Environmental Protection Agency. 1992. Consensus Method for Determining Groundwaters Under
the Direct Influence of Surface Water Using Microscopic Particulate Analysis (MPA). Manchester Environmental
Laboratory Report #910/9-92-029.53 p.
Vinyard, W.C. 1979. Diatoms of North America. Mad River Press, Inc., Eureka, CA.
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Appendix 1.
The microscope portion of this procedure depends upon very sophisticated optics. Without proper alignment and
adjustment of the microscope the instrument will not function at maximal efficiency and the probability of obtaining the
desired image will not be possible. Consequently, it is imperative that all portions of the microscope from the light
sources to the oculars are properly adjusted.
While microscopes from various vendors are configured somewhat differently, they all operate on the same general
principles. Therefore, slight deviations or adjustments may be required to make these guidelines work for the particular
instrument at hand.
1) Transmitted Light Adjustment. This section assumes that you have successfully replaced the transmitted
bulb in your particular lamp socket and reconnect the lamp socket to the lamp house. Make sure that you have
not touched any glass portion of the transmitted light bulb with your bare fingers while installing it. These
instructions also assume the condenser has been adjusted to produce Kohler illumination.
Step 1 Usually there is a diffuser lens between the lamp and the microscope which either must be removed
or swung out of the light path. Reattach the lamp house to the microscope.
Step 2 Using a prepared microscope slide and a 40 X objective, adjust the focus so the image in the oculars
is sharply defined.
Step 3 Without the ocular or Bertrand optics in place the pupil and filament image inside can be seen at the
bottom of the tube.
Step 4 Focus the lamp filament image with the appropriate adjustment on your lamp house.
Step 5 Similarly, center the lamp filament image within the pupil with the appropriate adjustment(s) on your
lamp house.
Step 6 Insert the diffuser lens into the light path between the transmitted lamp house and the microscope.
2) Adjustment of Interpupillary Distance and Oculars for Each Eye. These adjustments are necessary, so eye
strain is reduced to a minimum. These adjustments must be made for each individual using the microscope.
this section assumes the use of a binocular microscope.
A) Interpupillary Distance. The spacing between the eyes varies from person to person and must be
adjusted for each individual using the microscope.
Step 1. Place a prepared slide on the microscope stage, turn on the transmitted light, and focus the
specimen image using the coarse and fine adjustment knobs.
Step 2. Using both hands, adjust the oculars in and out until a single circle of light is observed while
looking through the two oculars with both eyes.
B) Ocular Adjustment for Each Eye. This section assumes a focusing ocular(s). This adjustment can
be made two ways, depending upon whether or not the microscope is capable of photomicrography
and whether it is equipped with a photographic frame which can be seen through the binoculars.
Precaution: Persons with astigmatic eyes should always wear their contact lenses or glasses when
using the microscope.
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Microscopic Particulate A nalysis (MPA) for Filtration Plant Optimization
1) For microscopes not capable of photomicrography. This section assumes only the right
ocular is capable of adjustment.
Step 1. Place a prepared slide on the microscope stage, turn on the transmitted light, and
focus the specimen image using the coarse and fine adjustment knobs.
Step 2. Place a card between the right ocular and eye keeping both eyes open. Using the
fine adjustment, focus the image for the left eye to its sharpest point.
Step 3. Now transfer the card to between the left eye and ocular. Without touching the
coarse or fine adjustment and with keeping both eyes open, bring the image for the
left eye into sharp focus by adjusting the ocular collar at the top of the ocular.
2) For microscopes capable of viewing a photographic frame through the viewing
binoculars. This section assumes both oculars are adjustable.
Step 1. Place a prepared slide on the microscope stage, turn on the transmitted light, and
focus the specimen image using the coarse and fine adjustment knobs.
Step 2. After activating the photographic frame, place a card between the right ocular and
eye keeping both eyes open. Using the correction (focusing) collar on the left
ocular focus the left ocular until the double lines in the center of the frame are as
sharply focused as possible.
Step 3. Now transfer the card to between the left eye and ocular. Again keeping both eyes
open, bring the image of the double lines in the center of the photographic frame
into as sharp a focus for the right eye as possible by adjusting the ocular correction
(focusing) collar at the top of the right ocular.
3) Calibration of an Ocular Micrometer (for Whipple grid also) This section assumes that
an ocular reticle has been installed in one of the ocular by a microscopy specialist and that
a stage micrometer is available for calibrating the ocular micrometer (reticle). Once installed
the ocular reticle should be left in place. The more an ocular is manipulated, the greater the
probability is for it to become contaminated with dust particles. This calibration should be
done for each objective in use on the microscope. If there is an optivar3 on the microscope,
then the calibration procedure must be done for the respective objective at each optivar
setting.
Step 1. Place the stage micrometer on the microscope stage, turn on the transmitted light,
and focus the micrometer image using the coarse and fine adjustment knobs for the
objective to be calibrated. Continue adjusting the focus on the stage micrometer so
you can distinguish between the large (0.1 mm) and the small (0.01 mm) divisions.
Step 2. Adjust the stage and ocular with the micrometer so the 0 line on the ocular is
exactly superimposed on the 0 line on the stage micrometer.
Step 3. Without changing the stage adjustment, find a point as distant as possible from the
two 0 lines where two other lines are exactly superimposed.
Registered trademark product of the Zeiss company. A device between the objectives and the oculars that is
capable of adjusting the total magnification.
23
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Microscopic Paniculate A nalysis (MPA) for Filtration Plant Optimization
Step 4. Determine the number of ocular micrometer spaces as well as the number of
millimeters on the stage micrometer between the two points of superimposition.
For example: Suppose 48 ocular micrometer space equal 0.6 mm.
Step 5. Calculate the number of mm/ocular micrometer space.
For example:
0.6mm
0.0125 mm
48 ocular micrometer spaces ocular micrometer space
Step 6. Since most measurements of microorganisms are given in um rather than mm, the
value calculated above must be converted to um by multiplying it by 1000 um/mm.
For example:
0.0125 mm >
Ocular Micrometer Space
1.000 um
12.5 um
mm Ocular Micrometer Space
Step 7. Follow steps 1 through 6 for each objective. It is helpful to record this information
in a tabular format, like the example, which can be kept near the microscope.
Item
#
1
2
3
4
Obj.
Power
10 X
20 X
40 X
100 X
Description
N.A.C =
N.A. =
N.A. =
N.A.=
No of
Ocular
Microm.
Spaces
No. of
Stage
Microm.
mm"
urn/Ocular
Micrometer
Space"
1000 um/mm
" (Stage Micrometer length in mm x (1000 um/mm) -r No. ocular micrometer spaces
c N.A. = Numerical aperture. The numerical aperture value is engraved on the barrel of the objective.
4) Kohler Illumination. This section assumes that Kohler illumination will be established for each DIC
or HMO objective. Each time the objective is changed, Kohler illumination must be reestablished for
the new objective lens. Previous sections have adjusted oculars and light sources. This section aligns
and focuses the light going through the condenser underneath the stage at the specimen to be observed.
If Kohler illumination is not properly established, then DIC or HMO woptics will not work to their
maximal potential. These steps need to become second nature and must be practiced regularly until
they are a matter of reflex rather than a chore.
Step 1. Place a prepared slide on the microscope stage, move the required objective into place, turn
on the transmitted light, focus the specimen image using the coarse and fine adjustment
knobs.
24
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
Step 2. At this point, both the radiant field diaphragm in the microscope base and the aperture
diaphragm in the condenser should be wide open. Now close down the radiant field
diaphragm in the microscope base until the lighted field is reduced to a small opening.
Step 3. Using the condenser centering screws on the front right and left of the condenser, move the
small lighted portion of the filed to the center of the visual field.
Step 4. Now look to see whether the leaves of the iris field diaphragm are sharply defined (focused)
or not. If they are not sharply defined, then they can be focused distinctly by changing the
height of the condenser up or down while you are looking through the binoculars. Once you
have accomplished the precise focusing of the iris field diaphragm leaves, open the radiant
field diaphragm until the leaves just disappear from view.
Step 5. The aperture diaphragm of the condenser is adjusted now to make it compatible with the total
numerical aperture of the optical system. This is done by removing an ocular, looking into
the tube at the rear focal plane of the objective, and stopping down the aperture diaphragm
iris leaves until they are visible just inside the rear plane of the objective.
Step 6. After completing the adjustment of the aperture diaphragm in the condenser, return the ocular
to its tube and proceed with the adjustments required to establish either DIG or HMO optics.
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Appendix 2.
Use of Electronic Particle Counter
Electronic Particle Counting for Filtration Plant Optimization
Introduction
Alternative methods for determining filtration efficiency, such as particle size analysis, have been recommended in the
"Guidance Manual for Compliance with Filtration and Disinfection Requirements for Public Water Systems Using
Surface Water Sources", USEPA, March 91. The data accumulated from particle size analysis should be used in
conjunction with microscopic participate analysis data to determine over-all filtration efficiency. The Federal Register,
Vol. 59 number 28, Feb. 10, 1994, states that particle counting data could be used as a tool for treatment process
efficiencies and could possibly be used as a surrogate for Giardia and Cryptosporidium monitoring. Either electronic
particle counters or treatment plant in-line installations that measure continuously can be used. These instruments give
particle size ranges and the number of particles per size range. A comparison of raw water particle counts verses the
finished water particle counts can be used to calculate percent removal or log reduction and an estimate of filtration
efficiency established.
Sample Collection
1.0 The samples collected for electronic particle counting can be grab samples or composite samples. If
a composite sample is collected, then the procedure used to collect and process the sample is
described in the MPA for Filtration Plant Optimization procedure. A subsample from this composite
sample will need to be diluted with particle free water before analysis. However, there are potential
problems with composite sampling that need research and depend greatly on the type of water being
analyzed.
2.0 Grab samples require little processing and may be useful for this reason. The main concern with the
grab sample is that collection occurs over a very discrete amount of time and therefore may not be
representative of the water supply. This problem can be resolved by either showing repeatability or
by comparing the grab sample results to a composite sample which is more representative.
3.0 Grab sampling: "dedicated" glass containers are to be used. A dedicated container is one which is
used exclusively for a certain type of water (one set for raw waters and another set for finished
waters). These containers should be cylindrical glass bottles that have been scrupulously cleaned with
a mild laboratory detergent and then rinsed a minimum of three times in particle-free water. Prior to
sampling, the container should be rinsed a minimum of three times with the water being sampled.
Plastic bottles should be avoided because they may shed particles into the sample.
4.0 Collection: Collect grab samples as close to the source as possible. A continuously flowing tap is
recommended, if not available, flush the sample tap for 5 minutes prior to sampling. Run the water
down the inside of the bottle to lessen air entrapment. Make sure to label the bottle: sample
identification, date, time, and sampler name.
5.0 Holding time: Analyze as soon as possible after collection. Raw samples are especially prone to
organic growth, adsorption of particles to the bottle walls and decay of original sample, all resulting
in alteration of the particles. Filtered, or otherwise treated samples, are not quite as critical, but should
be analyzed assoon as possible. If analysis can not happen immediately samples should be stored at
4°C and sealed with a teflon screw bottle cap. Do not expose the sample to sunlight or let it freeze.
26
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
Particle Analysis
6.0 Analysis by electronic counter for grab and composite samples: Electronic counting can be performed
by one of two types of counters; a light blockage device (HIAC, Met One, Hach or Particle
Monitoring System to name a few) or an electrical sensing zone device (Coulter Counter or Elzone).
7.0 Prior to analysis, verify the instrument's sizing capabilities according to manufacturer's instructions
using latex beads representing the particle sizes of interest. If a light blockage device is used, the
calibration is done during installation and on a routine basis recommended by the manufacturer. The
most widely accepted dimensions of Giardia cysts are 7-12 um. Cryptosporidium oocysts are in the
dimensional range of 3 - 7 um. Individual electronic particle counters measure these organisms
differently. Testing of individual instruments will be needed to determine the actual size measured
by a specific particle counter.
8.0 If sample has been refrigerated bring it to room temperature very slowly. Mix samples gently, by
swirling just prior to analysis. Minimize bubble formation, do not shake.
9.0 Run at least 3 rinse samples with particle free water to stabilize the instrument.
10.0 After running the sample in triplicate, run a rinse with particle free water or electrolyte to re-stabilize
the instrument.
11.0 Always run the cleanest sample first and proceed to the most concentrated.
12.0 Between analyses, keep particle-free solution in the instrument chamber. The sample is run in
triplicate to assess the instrument precision. The individual results should not vary more than 10 %
from the average of the three runs, except in low particle count water (less than 10 particles per mL).
13.0 To avoid coincidence errors, in concentrated samples (raw samples), dilution of the final sample will
undoubtedly be required. Coincidence occurs when more than one particle passes through the detector
at a time, causing inaccurate counting and diameter measurements. This dilution should be done with
the particle-free solution recommended by the manufacturer. Generally speaking, a dilution between
1:5 to 1:20 with particle-free solution will suffice. It is best to use a dilution as close to tolerance for
coincidence error as possible to decrease the number of background counts. It may also be necessary
to screen filter the sample if large debris repeatedly block the orifice tube. For composite samples
1:1000 dilutions are not uncommon. Any pipettes or glassware used to make dilutions should be
calibrated.
14.0 The average of the three values obtained are recorded in the corresponding box on the data sheet.
15.0 Calculation of the percent removal and log reductions can be done for both the total number of
particles or for each size range. See example data sheet.
27
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
References for Particle Counting:
1. Vasconcelos, GJ. and S.I. Harris. Procedure draft. Microscopic Particulate Analysis and Particle Size
Analysis for Determination of Filtration Efficiencies. EPA Manchester Environmental Laboratory.
2. A.P.H.A., A.W.W.A, W.E.F. 1995. Section 2560 Particle Counting and Size Distribution (Proposed) in: 19th
Edition Standard Methods, A.P.H.A., Washington, DC, pp 2-60.
3. Federal Register. 1994. Proposed Monitoring Requirements for Public Drinking Water Supplies:
Cryptosporidium, Giardia, Viruses, Disinfection Byproducts, Water Treatment Plant Data and Other
Information Requirements; Proposed Rule, February 10,1994. 40 CFR Part 141. Vol. 59, No. 28 Proposed
Rules, pp 6336.
4. Hargesheimer, E.E., C.M. Lewis, and C.M Yentsch 1992. Evaluation of Particle Counting as a Measure
of Treatment Plant Performance. A.W.W.A. Research Foundation Denver. pp 319.
5. USEPA. Science and Technology Branch. 1991. Guidance Manual for Compliance with the Filtration and
Disinfection Requirements for Public Water Systems using Surface Water Sources. Criteria and
Standards Division. Office of Drinking Water. Washington. D.C.
28
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Appendix 3
Sample Data Forms and Report Forms
Microscopic Participate and Particle Size Analysis
For Water Treatment Evaluations
Lab # Raw
Lab # Finished
Water Treatment Plant
Sample sites: Raw
Sampler's name
Agency
Finished
Raw water source: (circle one)
River
Infiltration gallery
Field Measurements:
Creek
Horizontal Collector
Spring well
Other
Raw
Finished
T. Cl
F. Cl
Turb
(MTU)
PH
Temp
(C)
TC/
100ml
FC/
100ml
Operational Parameters: circle appropriate choice
Pre-treatment
Filtration
Disinfection
alum
R. sand
Cl
lime
S.sand
Cl-A
Polymer
Press.
Filter
Ozone
carbon
Cartr
Other
other
other
Processing Information;
Total volume filtered (Liters)
Total filter sediment collected
(Packed pellet) (g)
Centrifugate volume/ 100 liters
Raw
Finished
29
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Particle Size Distribution and Percent Removal Using Microscope.
Average of 20 - 30 fields @ 100 X magnification
Particle Size category
< 10 urn
10 - 25 urn
25 - 100 um
100 - 200 um
>200um
Total
Number in
Raw Water
Number in
Finished Water
Percent
Removal
Particle Size Distribution and Percent Removal Using Electronic Particle Counter.
Channels
Number in
Raw Water
Number in
Finished Water
Percent
Removal
30
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Particle Counting Example
Filtration Efficiency
Using an Electronic Particle Counter*
Date:
Raw Water:
Finished Water:
May 20, 1994
Raw River Water
Finished Water
Samples:
Code number:
Code number:
Composite or Grab
2
1
Particle size (urn)**
2
3
5
10
12
>15
Total
Raw Particles/mL **
40,556.3
8,225.6
20,985.5
12,358.8
4,569.8
2345.6
89,041.6
Finished Particles/ml
4563
100
125
96.6
5.2
8
791
% Reduction
98.8749 %
98.7843
99.4044
99.2184
99.8862
99.6589
99.1115
Log Reduction
1.9488
1.9152
2.225
2.107
2.9439
2.4672
2.0514
* All limitations of the analytical methods, laboratory dilutions and instrument apply.
** Measured in equivalent spherical diameter.
**'Numbers represent the average from 3 subsample counts.
Raw vs. Finished
10 12
Particle Size (urn)
Total
Raw Particles/ml*
Finished Particles/ml***
31
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
Analysis for Waterborne Participates
Customer
PWSID*
Sample Information:
Date/Start: Mrs Date/Stop: Mrs Sampler:
Gallons: Filter Colon Centrifugate: tnL/100 gals
Results of Microscopic Particulate Analysis':
Amorphous Debris: /uM diameter
Nondiatomaceous Algae:
Diatoms:
Plant Debris:
Rotifers:
Nematodes:
Pollen:
Ameba:
Ciliates:
Coloriess Flagellates:
Crustaceans:
Other Arthropods:
Othen
Comments:
Type of Wash water used; total or natural count used, type of counting chamber used
Laboratory Information:
; ; Mrs; ; ;
Results submitted by:
32
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
Appendix 4
Formulation of McFarland Standards
McFarland standards provide laboratory guidnace for the standardization of numbers of bacteria for susceptibility testing
or other procedures requiring a standardization of inoculum. They are devised to replace the counting of individual cells
are are designed to correspond to appropriximate cell densities.
1) Make solution 1 1 % H,SO4
Make solution 2 1.175 % BaCl2
Dispense in the following amounts of desired standard totaling 10 ml in 16 X 125 mm tubes. Cap and label. Should be
replaced every six months.
Standard Concentration
0.5
1
2
3
4
5
BaCU Volume in ml
0.05 ml
O.lml
0.2ml
0.3ml
0.4ml
0.5 ml
H,SO4 volume in ml
9.95 ml
9.9 ml
9.8ml
9.7ml
9.6ml
9.5ml
33
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Microscopic Partieulate A nalysis (MPA) for Filtration Plant Optimization
Appendix 5
Figures for Document
Figure 1
Raw Water Sampling Apparatus
pressure gauge
backflow preventer
>- - WATER SOURCE
pressure regulator
Inlet hose T
I (pump optional at these points)
quick connects \
v
filter
holder
34
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Figure 2 Finished Water Sampling Apparatus
(For chlorinated water only)
pressure regulator
backflow preventer
«^ . WATER SOURCE
| (pump optional at these points)
quick connects \
V
proportloner
(for disinfected water)
35
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Microscopic Paniculate Analysis (MPA) for Filtration Plant Optimization
Figure 3
Processing Flow Chart for Palmer Maloney Counting Chamber
EPA sampler
10p.s.i.
Minimum of 100 liters raw;
1,000 liters finished
Paniculate extraction
Handwash
I
Mechanical wash
(Stomacher* 3500)
Record volume
of total paniculate solution in liters
Thoroughly mix
solution
Transfer 200 ml to 250 ml conical
centrifuge bottle
Vortex 15 sec.
Withdraw 0.10 ml, inoculate PMCC
Examine at lOOx magnification minimum
plankters/field
Microscopic examination
(100x phase contrast, DIC, HMO)
plankters/field
Microbiota
TNTC
Microbiota nor
TNTC
T
Use Whipple grid
@400x
magnification
i
Total of 100
organisms are
counted
Entire
PMCC
counted
^
w
plankters/field
Centrifuge 10min.
@1050g
aspirate to pellet
Withdraw 0.1 ml
inoculate PMCC
< 10 plankters/field
Additional 200 ml subsample
Count particles in size
ranges
(<10,10-25, 25-100,
100-200, >200)
in 20-30 random
Whipple grid fields
on PMCC
36
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Microscopic Paniculate A nalysis (MPA) for Filtration Plant Optimization
Figure 4 Centrifugate Pellet Volume Determination
Pellet Centrifugate Measurement
±
Remove aliquot equivalent to 1001.
Place in Conical Centrifuge Tubes
Centrifuge 10 min. @ 1050g
Aspirate supernatant
Using calibrated tube set; measure
pellet volume
Record Centrifugate pellet volume
37
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Microscopic Particulate Analysis (MPA) for Filtration Plant Optimization
Figure 5 Scanning Cover Slip Method
-------
pic Paniculate Analysts (MPA) for Filtration Plant Optimization
Appendix 6
Example of Sample Calculation
Assume that a 1000 Liter (380 gal) water sample was collected. The sample was eluted resulting in 3 L of
paniculate solution. The solution was thoroughly mixed and a 200 mL aliquot was placed in a centrifuge bottle. A
0.1 mL aliquot from the centrifuge bottle was examined in a Palmer-Maloney Chamber but only 5 plankters per field were
observed, so the 200 mL subsample was centrifuged and 150 mL of the supernatant was aspirated. The remaining 50
mL of supernatant and pellet was thoroughly mixed before 0.1 mL was examined in a Palmer-Maloney Chamber.
Because 20 plankters per field were observed, the plankters were identified and counted.
The liter equivalent-in the centrifuge bottle was calculated based upon these facts:
3000 mL of particulate solution = 200 mL aliquot in centrifuge bottle
1000 L sampled 66.667 Liter equivalent in centrifuge bottle
The liter equivalent in the first 0.1 mL subsample (x), where 5 plankters per field were observed, is calculated
using the proportion ratio inJQJLL., -
r*"~ *
200 mL in centrifuge bottle = 0.1 mL [observed in Palmer-Maloney Chamber]
66.667 Liter equivalent in centrifuge bottle x = 0.033 Liter
i i
The liter equivalent of the remaining 50 mL of supernatant and pellet is 66.634 which is the 66.667 Liter
equivalent in the centrifuge bottle.prior to centrifugation minus the 0.033 L equivalent withdrawn in the first 0.1 mL
subsample (calculated in 10.3.1).
The following organisms were identified and counted in the second O.lmL subsample when the entire Palmer-
Maloney Cell is scanned at lOOx: r-
10 Keratella (rotifers)
3 Vonicella (ciliates)
1 Nematode ..M.. -*.
The numbers per 100 Liter are calculated using the proportion ratios in 11.1.5.1 and 11.1.5.2:
10 Keratella = x = 5.000
O.lmL 50 mL
Because the 50 mL of paniculate solution in the centrifuge bottle is equivalent to 66.634 L:
5.000 = 7503 Keratella
66.634 L equivalent 100 L
Similarly,
3 Vonicella = 1500
O.lmL 50 mL
1500 = 2254 Vonicella
66.634 L 100 L
39
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Microscopic Participate Analysis (MPA) for Ftitratiom:ei**Qlttimiatitm
and
1 Nematode =
0.1 mL 50 mL
500 = TSONematodes
66.634 L 10ft L
The algae were too numerous to count at lOOx so 100 algal cells were counted in 5 whipple grid fields at 400x.
The number of algae is calculated as in 11.2.3,11.2.4, and 11.2.5:
100 alga counted x 1000mm * No. algae/mJL
A x 0.4 x 5 fields
A = the area of the whipple grid field, which must t» calculated for the microscope, used for the counts (section
12.0). For example A = .074. (0.4 is the depth of the Palmer-Maloney chamber.)
then,
100 x 1000 = 675,67y.67algrtcells/oiL
.074 x .4 x 5
then,
675.675.67 * 33.783,783
mL50 mLs
and because 50 mL is equivalent to 66.634 L:
33.387.783 = 50.700300 Algae
66.634L 100 Liters
Report the organism values in significant figures using the guidelines in Standard Methods. For this qxajnple the values
would be reported in #'s/100 L as follows: 8,000 Keratella
2,000 Vorticella
800 Nematodes
50,000,000 Algae
40
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