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Introduction to Lake Surveys: Laboratory Techniques

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1 Introduction to Lake Surveys: Laboratory Techniques
Unit 3: Module 9

2 Objectives Students will be able to:
define alkalinity and hardness in water. identify methods used to measure and analyze the alkalinity and hardness in water samples. identify methods used to determine the amount of specific nutrients in water. interpret data from nutrient standard calibration curves. explain methods used to measure total suspended solids in water samples. calculate the total suspended solids in water samples. explain methods used to measure turbidity. evaluate and compare turbidity data against specified standards.

3 Objectives cont. Students will be able to:
describe procedures used for determining biochemical oxygen demand. explain methods used to determine algal biomass and biovolume. compare and contrast spectrophotometers and fluorometers. identify methods used to measure algal chlorophyll. estimate the biomass and biovolume for periphyton samples. describe procedures used to measure bacterial colonies in water samples. determine methods used to measure biomass of aquatic vegetation. identify methods used to measure benthic invertebrates and zooplankton. analyze the properties of benthic sediments.

4 Basic water quality assessment – lab
Goals – lectures and labs focus on analyzing samples in lake surveys and on parameters used in lab experiments Water chemistry – alkalinity and hardness nutrients by colorimetry and kits suspended sediments (TSS) turbidity organic matter (BOD), color The Module 9 labs will be: A set of detailed lab manual type water chemistry “recipes” (as PDF files) pH and alkalinity Nutrients by colorimetry With spectrophotometer Via HACH type kits TSS and turbidity Organic matter (BOD5) and color Dissolved oxygen (Winkler titration) and BOD Linked to sediment O2-demand (respiration) and Fish Respiration lab in WOW I Chlorophyll a for both phytoplankton and periphyton Algal enumeration and biomass estimates Zooplankton and benthos community assessments Aquatic vegetation Bacteria; fecal coliforms and E. coli. References for key texts and permanent website manuals (EPA, USGS, APHA, etc) Lab experiments, where possible, that use techniques to illustrate an ecological process such as: DO/BOD for comparing plankton to fish to benthic respiration (as per a WOW I lab) Road salt lab from WOW I TSS lab from 2002 WOW 2 exercise

5 Basic aquatic community assessment
Algae and bacteria (chlorophyll-a, microscopy, plate counts) Aquatic vegetation and attached algae (periphyton) Zooplankton Sediment bulk properties Benthic organisms Microbial pathogen indicators Fecal coliforms and E. coli

6 Alkalinity and hardness
Photo of pH test References: USGS TWRI Book 9 Chapter 6.6 Alkalinity and Acid Neutralizing Capacity:http://water.usgs.gov/owq/FieldManual/Chapter6/section6.6/ APHA Standard methods for the examination of water and wastewater. American Public Health Association, Washington, D.C. Lind, O Handbook of common methods in limnology 2nd Ed. Kendall/Hunt Publishing, Co. Dubuque, IA.

7 Alkalinity and hardness - what is it?
Alkalinity: a measure of the ability of a water sample to neutralize strong acid Expressed as mg CaCO3 per liter or microequivalents Alkalinities in natural waters usually range from 20 to 200 mg/L Hardness: a measure of the total concentration of calcium and magnesium ions Expressed as mg CaCO3 per liter The USGS (USGS 2001) defines alkalinity as ‘the capacity of solutes in an aqueous system to neutralize acid” and ANC as “the capacity of solutes plus particulates in an aqueous system to neutralize acid”. Following the 1980’s acid rain surveys the term ANC or acid neutralizing capacity has come into use. In most cases alkalinity and ANC are virtually synonymous. Therefore, strictly speaking: ANC determined on raw, unfiltered samples Alkalinity determined on filtered samples Both are determined by titration with a dilute, strong acid (usually 0.02 – 0.1 N H2SO4) to an endpoint of 4.5 – 4.2. Alkalinity and hardness classifications : see Modules 2 - 3 Hardness, in the past, was measured as the capacity of water to precipitate soap when a liquid soap solution was shaken with the water sample to form a lather persisting for 5 minutes (Lind 1985).

8 Alkalinity and hardness - how to sample
Usually collected at the surface in lakes (0 to 1m depth) Keep the sample cool (4oC refrigerated) and out of direct sunlight Eliminating the head space, or air within the container, prevents the exchange of gases between the sample and any air inside the container. EPA recommended holding time is 24 hrs at 4 oC with NO headspace Why just surface sampling ? Only because in most cases the differences between lakes is far greater than the differences between depths in the same lake. We’re usually most interested in classifying lakes into softwater versus hardwater groupings.

9 Alkalinity and hardness- why measure?
The alkalinity of natural waters is usually due to weak acid anions that can accept and neutralize protons (mostly bicarbonate and carbonate in natural waters). Usually expressed in units of calcium carbonate (CaCO3) The ions, Ca and Mg, that constitute hardness are necessary for normal plant and animal growth and survival. Hardness may affect the tolerance of fish to toxic metals. Wetzel, R. and G. Likens Limnological Analyses, 3rd edition. Springer-Verlag, New York, Inc. APHA Standard methods for the examination of water and wastewater. American Public Health Association, Washington, D.C. Harder water renders many heavy metals less toxic. Originally, hardness was understood to be a measure of the capacity of water to precipitate soap and it is the magnesium and calcium ions present in the water that do this. Total hardness is defined as the sum of the Ca and Mg concentrations (APHA 1998). Note: The units “mg CaCO3” is rather strange. This is an historical relict from days when alkalinity was predominately measured by drinking water analysts. They were most concerned about maintaining a moderate level of carbonate/bicarbonate alkalinity in the water. Too much leads to excessive marl, the white precipitate that forms in home plumbing systems and poor soap lathering. Too little leads to excessive pipe corrosion.

10 Alkalinity – analysis pH meter Buret* Thermometer
Magnetic stirrer and stir bar Top loading balance We also place a wood block beneath the beaker that prevents the stir plate from heating the sample. A buret is a long, graduated glass tube with a tapered tip like a pipet and a valve that is opened to allow the reagent to drip out of the tube. The amount of reagent used is calculated by subtracting the original volume in the buret from the volume left after the endpoint has been reached. Alkalinity is calculated based on the amount used. Titrators forcefully expel the reagent by using a manual or mechanical plunger. The amount of reagent used is calculated by subtracting the original volume in the titrator from the volume left after the endpoint has been reached. Alkalinity is then calculated based on the amount used or is read directly from the titrator. Digital titrators have counters that display numbers. A plunger is forced into a cartridge containing the reagent by turning a knob on the titrator. As the knob turns, the counter changes in proportion to the amount of reagent used. Alkalinity is then calculated based on the amount used. Digital titrators cost approximately $90. Digital titrators and burets allow for much more precision and uniformity in the amount of titrant that is used. A microburet is shown in the image above. It dispenses up to 2 mL of titrant with greater than or equal to 0.003 mL accuracy.

11 Alkalinity- analysis Reagents
0.04 N H2SO4 (see method for details on preparation) Total alkalinity analysis involves titration until the sample reaches a certain pH (known as an endpoint) At the endpoint pH, all the alkaline compounds in the sample are "used up" The amount of acid used corresponds to the total alkalinity of the sample The result is reported as milligrams per liter of calcium carbonate (mg/L CaCO3) The value may also be reported in milliequivalents by dividing by 50 0.04N H2SO4 titrant: Dilute 2.27 mL concentrated H2SO4 [*C] to 2 liters with Milli-Q water. Standardize against a 100 eq /L THAM (Tris hydroxymethyl aminomethane) solution prepared by dissolving g dried THAM in 100 m/L Milli-Q water in a 1000m/L volumetric flask. Dilute to the mark with MQW. See method (PDF document ) for details. Different endpoints are used for different ranges of alkalinity. See Table 1 in Alkalinity method (PDF) for suggested equivalency points. Minor errors that may result from drift in pH readings are checked using a dilute sulfuric acid solution giving a low pH that is considered relatively stable (low ionic strength buffer or LISB). The commercial buffers used to calibrate probes are of very high ionic strength, much higher than most natural surface waters. A LISB, a dilute sulfuric acid solution of pH around 3.7 can be used to determine if a pH probe is capable of responding to low ionic conditions. Divide by 50 ? The formula weight of CaCO3 divided by its valence of 2.

12 Alkalinity- analysis or Where:
B = mL titrant first recorded pH (i.e., to pH = 4.5) C = total mL titrant to reach pH 0.3 unit lower, and N = normality of acid (titrant) Alkalinity as mg CaCO3/L can be converted to meg/L by dividing by 50.

13 Hardness – analysis Hardness, in units of mg CaCO3/L
Hardness is, ideally, determined by calculation from the separate determinations of calcium and magnesium. Hardness, in units of mg CaCO3/L Where Ca and Mg are in mg/L Calcium and magnesium are determined separately using an atomic absorption (AA) spectrometer, APHA 3111 B or by ion chromatography. There is also an EDTA titrametric method (APHA 2340C) which provides an estimate of the combined hardness due to calcium and magnesium.

14 Alkalinity and hardness – analysis
There are also titration test kits available for both alkalinity and hardness Hach Chemical Co (www.hach.com) and LaMotte Chemistry (www.lamotte.com) are two companies that market easy to use test kits.

15 Nutrients WOW figure

16 Nutrients: colorimetry & spectrophotometry
Overview of the colorimetric analysis of the nutrients nitrogen and phosphorus using spectrophotometry Specific techniques for students to review in or out of class included: developing calibration curves QA/QC : standards, spikes, etc. Teachers: Some recurring themes to consider harping on – Note: There will be accompanying information in the form of: - .pdf and .doc files of lab handout(s) - .pdf and .doc files with detailed analytical protocols Units: ug/L = 1000 ppb = 1 ppm = 1 mg/L (aquatic scientists must learn to beware of units changing “form” suddenly) Magnitude of water chemistry concentration: Major ions are usually in ppm Nutrients and micronutrients are usually in ppb (although many State and Federal databases use mg/L (i.e., ppm ) almost exclusively Context is often important: What defines a pollutant usually relates to the so-called beneficial use of the waters. WHAT IS WATER QUALITY ? - Are you concerned with human health or environmental health? - Do you need the water to be fishable and swimmable or do you need it to be drinkable? Ex. #1: Lake phosphorus levels: 100 parts per billion (ppb) is considered eutrophic (usually undesirable) Cola phosphorus: 1000s of parts per billion is considered tasty! Drinking water: There is no limit to the amount of phosphorus allowed (Phosphorus, in fact, may be added at parts per million to retard lead (Pb) leaching from house or City pipes) Ex. #2: Lake nitrogen: >500 ppb is considered high Drinking water: Considered OK if <10,000 ppb (as nitrate-N) Ex. #3: Lake mercury (Hg): >6.9 parts per trillion (ppt) = high ecological risk Drinking water: OK if <2,000,000 ppt Ex. #4 User perceptions may vary regionally as well regarding what lake water transparency is acceptable. This means that a total phosphorus level of 50 ppb P that results in a Secchi transparency of only 1.5 m in Iowa ponds may be very acceptable in a region where are no clear, blue lakes like Lake Tahoe. They might still be good for fishing, boating, and even swimming. But in NE Minnesota, this much total phosphorus and its associated Secchi transparency readings is perceived as very poor water quality. In this neck of the woods people want >4 m Secchi readings, which may translate to total phosphorus values of <10-15 ppb P.

17 Nutrients - how to sample
Usually collected from discrete depths Keep samples cool and dark Freeze unless you can run in <24 hrs Follow APHA recommendations See Module 8 Part C for more details. Some total phosphorus protocols call for acid addition as a preservative if the sample is only to be analyzed for TP. To do this add 1 mL concentrated HCl per liter of sample. USEPA recommended holding times at 4o C are as follows: Ortho-P: 48 hrs if filtered TP: 24 hours NO3/NO2 : 48 hrs NH4: must be acidified to pH < 2 with H2SO4 APHA 1998 recommends: If the samples are to be frozen there is an indefinite holding time although it is prudent to run them within 30 days. Freezer space and the need to examine the data are usually the driving factors in getting the samples analyzed.

18 Nutrients: sample processing
Total phosphorus (TP) and total nitrogen (TN) analyses are made with whole, or raw, water Unfiltered sample Dissolved (soluble) fractions are with a filtrate Includes ortho-P, ammonium, nitrate and nitrite EPA and most states require the use of a membrane filter with a nominal pore size of 0.45 um most researchers use glass fiber filters Researchers however, often use glass fiber filters such as those used for TSS. In practice, it makes little difference.

19 Nutrients: colorimetry & spectrophotometry
Principles: Higher concentration of color = higher absorbance, as measured by a spectrophotometer add a dye that binds specifically to nutrient of interest measure the increase in “color” as an estimate of analyte concentration Prepare calibration standards - solutions with a range of nutrient concentrations Compare sample absorbances to calibration standard absorbances to estimate sample concentrations Principles: There is a detailed discussion of the Lambert - Bouguer Law of Absorption and of Beer’s Law in the textbook: Wetzel, R.G. and G.E. Likens Limnological Analyses. 3rd Edition. Appendix 3. Springer. The two laws can be combined to form what is often called Lambert-Beer’s Law which states that there is a linear (straight line) relationship between the absorbance of a solution and the concentration of a solute. This of course assumes that the solute is causing almost all of the light attenuation. In practice, clever analytical chemists have figured out how to get certain colored dyes to form from a chemical complex that is created when reagents bind specifically to a particular nutrient such as phosphate (PO4-4), ammonium (NH4+), or nitrate (NO3-). Other reagents are added to the water sample to create the proper chemical environment for the reaction to occur (such as a certain pH or ionic strength). The more of the nutrient present in the water, the more color is produced, so the intensity of a beam of light shining through a small tube of the stuff will be reduced. To quantify the actual concentration, a set of calibration standards are prepared very carefully that are analyzed identically to the lake or stream water samples. Based on the relationship generated by plotting the measured absorbances of the known standard concentrations, you generate a standard curve. From this you estimate the concentration in your sample (aka, the unknown). Notes: The Lambert-Beer “Law” is also used to characterize the light attenuating characteristics of the water column in a lake, wetland or stream. You will see this quantified as: Iz = Io e-kz where Iz = the intensity of light at a depth z, Io = initial light intensity at the water surface (actually just below the surface), z = the depth at which measurements of light intensity are made, and k = an extinction coefficient that is based on the assumed exponential attenuation of light with depth. Limnologists use various light sensors that are lowered to provide plots of different bands of light or specific wavelengths versus depth. Using some simple algebra, one can rearrange the equation to show that a plot of the natural logarithm of intensity versus depth should be linear. The slope of this line = the negative of the extinction coefficient. By either “eyeballing” the best straight line fit, or using a linear regression analysis from a hand calculator or spreadsheet software (Excel, QuattroPro or others), you can calculate the “best fit” extinction coefficient. Clearer lakes have smaller extinction coefficients than more turbid (cloudy) lakes. See Modules 2 and 3 and the Lake Ecology Primer (http://wow.nrri.umn.edu/wow/under/primer/page4.html) for further information. Also- light “behaves” differently depending on wavelength. The extreme ends of the visible spectrum (the reds and the violets) are absorbed more strongly than the green and blue wavelengths. Ultraviolet and infrared radiation are both more rapidly removed or attenuated than “middle” wavelengths. UV is also particularly well absorbed by the dissolved organic carbon compounds in water – the stuff that gives dark water lakes and streams their orange-brown appearance. In dark water systems, all of the UV may be absorbed in the upper 5-10 cms of the water column, in comparison to very clear water systems where it may be measurable and produce biological effects down to several meters in depth or more.

20 Nutrients: colorimetry & spectrophotometry
Add reagents to develop color Compare using a chart or color wheel using a colorimeter determining the absorbance using a spectrophotometer Low ….…. to ……. High Phosphate concentration Measuring the “color:” Simplest = Visual comparison to a chart with different shades. Often this is a disc inserted into a color wheel with a chamber that holds a vial or small test tube of the sample on one side and a chamber next to it that allows you to vary the “standards” until you get the best match. Often called a color “comparator.” Colorimeter: This is an instrument consisting of 1) a light source such as a tungsten-filament bulb; 2) a lens that focuses the light; 3) a colored filter that transmits light of the color which absorbed by the treated sample; 4) a sample compartment that holds a vial, test tube or cuvette of the sample; 5) a light sensor, like the one on your camera; and 6) electronics to convert the sensor signal to a meter or numerical digital readout – usually directly in concentration units such as parts per billion (ppb, also micrograms/liter = µg/L) or parts per million (ppm, also milligrams/liter= mg/L). Spectrophotometer: This instrument is basically an advanced version of a colorimeter. Instead of using a filter to select the right color for the test, it uses a prism or diffraction grating to select a narrow band of the desired wavelengths of light. The wavelength can be precisely tuned by rotating the prism or grating. A high quality “spec” with a bandwidth of ~ 2 nanometers, or nm, (meaning its accuracy for red light at 640 nm is about + 1 nm ) will cost several thousand dollars or more.

21 Color comparators and colorimetry
Test Kits – There are many brands available Images from Color Tube Color Disc Pocket Colorimeter Kits are also available from LaMotte Here are some typical kits and a colorimeter. A web search for “Water Chemistry Testing” will turn up many manufacturers. Hach Chemical Company is participating in Water on the Web via representation on our National Advisory Team. The Team meets annually to provide critical review of the project (http://wow.nrri.umn.edu/wow/overview/advisory.html)

22 Color measuring instruments
Bausch & Lomb spectrophotometer 20 Hach DR2400 portable spectrophotometer Spectrophotometry: Most spectrophotometers (usually just called “specs”) report two types of measurements: percent transmittance (%T) and absorbance (A or ABS). Percent transmittance is the ratio of the intensity of the light passing through the sample to the intensity of the light shining on the sample multiplied by 100% %T = 100 * (I/Io) Absorbance is the log10 of the transmittance: A = log10 (Io/I) = -logT Spectrophotometers can measure absorbance and transmittance over a range of wavelengths and you must select a wavelength and calibrate the instrument at that wavelength before making any measurements. Typically, the instruction manual will guide you through this process. In general the cuvette that holds the sample will be filled with deionized water (DIW) while reading on the %T function. A gain adjustment is tweaked to set it to exactly 100%, indicating maximum transmittance, and then the function is switched to ABSORBANCE. Another adjustment is made to set ABS = for 100%T by moving back and forth between the two. Website Links: As of this draft module, there was available a great set of detailed instructions about the workhorse Spec 20 that has been a feature of high school and university labs everywhere for decades. Go to The instrument was produced originally by Bausch & Lomb, Inc. and has gone through several cosmetic metamorphoses over the years. A simple web search for “Spec 20” will net you dozens of detailed instructions with excellent images that were assembled by chemistry and biology professors for classroom use. Cuvette notes: Handle carefully to avoid scratching them. Some are made of quartz, not glass, and may cost hundreds of dollars. Wipe all water and condensation off them (to prevent light interference). Be especially careful about pipetting directly into the cuvette. Remove it from the instrument first, since most water chemistry reagents are corrosive, being either acidic or basic or are solvent based (chlorophyll is usually extracted in 90% acetone or methanol). Wavelength choice: Wavelength (lambda) is selected to obtain the maximum sensitivity – i.e., the biggest change in absorbance per part per billion concentration change while minimizing interferences that might decrease absolute accuracy. These might be caused by compounds that also bind to the dye (arsenic at high levels can appear as a significant fraction of what you think is just ortho-P), or just by the color they add to the water. The dark water found in brown water lakes, streams and wetlands is an example. In some cases, fine particulates may add turbidity to a sample, causing erroneous absorbance readings. In practice, this usually means using wavelengths > 500 nm to minimize scattering effects that are more pronounced for “bluer” light. Phosphate is usually measured at 880 nm (infrared) or 675 nm (red); ammonium at 640 nm; and nitrate at 543 nm (actually we measure nitrite after chemically reducing the nitrate to nitrite). Using the wrong wavelength won’t trash your data – it will typically just decrease the sensitivity of the test.

23 Calibration standards
Standards are made from a concentrated stock solution that is precisely diluted to create “working standards” that are used and then discarded Calibration curves 3-5 concentrations spanning the expected range of the samples. always include a “blank” (= 0 ppb) of high purity deionized water (DIW). The better your DIW, the lower your detection limit usually if your glasssware is clean, your technique is good, and your reagents are clean. Usually there are about samples in a batch with another standards and quality assurance samples. Color development times vary according to the specific analyte, but all will usually continue to darken a bit after the allotted time. It may take several minutes to add reagents at each step to the samples and minutes to read the sample absorbances for a full batch after the color has developed. Therefore, it’s a good idea to set up duplicate sets of standards and arrange the tubes as a set of standards, then the samples, then the second set of standards and to read them in this order. The standards are then all pooled and included in the calibration curve (see details in subsequent slides). This allows some degree of control over sample colors that might change during the time that it takes to read their absorbances. When catastrophe strikes, such as when there’s a problem with the spec that takes an hour to solve, put the batch in a dark drawer and hope for the best. Samples are “precious” because you can’t get back out there to re-sample, and so you really rely on your standards and QA spikes to salvage the data. Ortho-P: Use dried KH2PO4, K2HPO4, NaH2PO4 or Na2HPO4 NH4-N and NO3-N: Use dried NH4NO3 as a dual standard (50% of each form)

24 Water chemistry “101” Procedure: See specific analyses
Reagents are added to each sample and standard identically Mix after each step Incubate at room temp or in water bath for 20 min to ~ 2 hrs, depending on the analyte Specific water chemistry procedures for ortho-P and ammonium-N are downloadable from the Water on the Web website at These two methods were selected because their color development time is relatively rapid – about 20 min for ortho-P and about 45 min for the UV ammonium-N method which allows both to be used (with some degree of luck and a good deal of preparation beforehand) in a 2-3 hour lab. A decent spectrophotometer (a Spec 20 will work) is needed, as well as some tubes, pipettes and glassware. We suggest a pre-made set of standards that can be included along with student-generated standards and water samples. The students can do the spikes for quality assurance (QA) as well. See Unit ___ Module ___ for further information about QA/QC.

25 Standard calibration curves
Good straight line fit: ABS = a + b*[Conc] NH4-N standards Range of NH4-N is 0 – 1000 (the bluest) ppb (= ug N/L) Range of PO4-P is also about 1000 ppb (shades of gray-blue) Notes: The concentrations of the standards are plotted as the independent variable (x-axis), because these were prepared. The dependent variable is Absorbance because it depends upon (is a function of) the standard concentration, which was determined by dissolving weighed amounts of high purity nutrient salts into high purity DIW using volumetric glassware and pipettes. Students should graph the data on graph paper and using spreadsheet software. Students should be shown how to use a hand calculator and spreadsheet software to perform a linear regression analysis to obtain the slope of the line (b), the y-intercept (a), and the regression coefficient (r2), which is essentially an estimator of the goodness of fit. Spreadsheet software typically also produces a P-value that represents the probability, or significance level, of the slope being greater than zero. Any calculator with two-variable statistics capability will usually perform a regression fit. R2 also estimates the fraction of the variability in the data that is explained by the fitted straight line. The r2 should typically be >0.99. Caution – Calculators often give you an r value that you have to square yourself. “R” is the correlation coefficient.

26 Estimating concentrations
So, if sample #3 had an absorbance of 0.290… Its concentration would be ~ 0.33 ppm N …

27 Standard curves – troubleshooting
#1 Example #1 – Live with it or re-run the batch Errors in preparing the 0.25 and 0.50 ppm standards perhaps ? #2 Example #2 – Fit a straight line from and a 2nd line from ugN/L Use non-linear quadratic instead of a line for ugN/L Re-read in smaller cuvette or dilute and re-run Errors in standard curves Variability in making standards – practice, practice, practice! Your pipetting technique is not consistent usually; or a particular standard may have been contaminated. It’s a good idea to make each standard concentration individually by diluting from an intermediate stock solution. Always make duplicates of each standard concentration – making each one individually. One common method is to maintain a concentrated stock of say 50 mg P/L (ortho-P) in a glass bottle in the refrigerator. On analysis day, it is diluted as follows: 2 mL: 100 mL DIW to form a working stock of 1 mg/L P (=1000 ug P/L). Then a set of standards can be prepared from the working stock as: 1 mL:100 mL DIW = 10 ug P/L 2.5 mL:100 mL DIW = 25 ug P/L 5 mL:100 mL DIW = 50 ug P/L 10 mL:100 mL DIW = 100 ug P/L 25 mL:100 mL DIW = 250 ug P/L 50 mL:100 mL DIW = 500 ug P/L NOTE: Since each set is prepared independently, there’s less chance of propagating a dilution error such as would occur if the 1000 ug/L stock was diluted 1:1 to get a 500 ug/L standard, which was then diluted 1:1 to get a 250 ug/L standard, etc. In this case an error in the first dilution carries over to subsequent sets of standards. Systematic errors can be caused by impure reagent used to make the primary stock an inaccurate balance, weighing error, or incorrectly using a pipette. These are all rather sinister errors because they are not revealed until you analyze someone else’s “CERTIFIED” standard. You can be very precise and have excellent repeatability, but be way off in absolute accuracy. Purchase commercially certified standards and make sure any lab work you contract out only goes to a state-certified lab. Most states now have programs to certify labs. However, even if certified, you should always check on the lab by talking to past clients and by asking to examine their quality assurance and quality control procedures. More on this in UNIT 4, Module 13. Labs also will run a Quality Control Check Standard (QCCS), along with a batch of samples for many analytes. This is usually a big batch of water that is either conservative (it won’t change over time, such as chloride) or has been preserved in some fashion to keep it constant (usually acidifying it to pH 2 with high purity acid or by adding some nasty toxicant). You then hope to obtain a value of ~ % of your original measured value, which you based on a whole bunch of replicates. A “good” value adds additional assurance to the quality of a particular batch of samples. Precision is estimated by doing replicates and calculating either: Coefficients of variation (CV) for n > 3 where CV (in %) = 100 * [standard deviation/mean]. A CV < % is desirable. As you get closer to the method’s level of detection (LOD) , the CV will usually rise considerably. For duplicates, the relative percent difference is used. For replicates x1 and x2, RPD = [x1-x2]/[2*mean]. Again, a value of < 10-15% is desirable. It’s usually expressed as an absolute value, since there is no difference between the replicates – you’re just looking for a measure of how different they are from their mean value. Spike Recovery (%R) – this is a particularly important part of QA/QC and essentially tests to ensure that there are no interfering compounds in the environmental water sample that make it respond differently to the water chemistry reagents than the deionized water (DIW) that is the matrix for your calibration standards. You enrich (spike) a particular sample that is being run independently, with a known amount of analyte and then run this in the same batch with your sample. %R = 100* [(spiked sample value)-(sample alone value)/known spike value] %R = 100* [(Xf -Xo)/S , where Xo = initial sample concentration, Xf = final sample concentration, and S = spike concentration Example – - Sample #3 was measured to be 330 ugN/L of ammonium-N (also written as 330 ug NH4-N/L) - We also spiked it separately with exactly 50 ugN/L of ammonium-N - In the same batch as #3, we measured #3-Spike to be 377 ugN/L - The %R = 100 * [( )/50] = 94%. - The accepted criterion for “passing” is usually % but may be tighter or looser depending on the project needs. Record Keeping: Labs should have running plots of their chemistry parameters from batch to batch. These include, calibration curve regression coefficients, r2, CV or RPD for replicates, %QCCS, and %R. Often the blank (0 ppb standard) absorbance is included. Together, these help you discover problems from bad reagents, bad lab DIW, or bad technique before you screw up more than a batch or two. The line becomes non-linear after ABS ~ 1.0 (~ 1000 ugN/L)

28 Some data from northern Minnesota lakes
Calibration curve = std ABS = ( ) + ( )* P R2 = n=12 Sample #1 = 11.2 ugP/L Sample #1 - Replicate = 12.6 ugP/L Sample # Spike = 59.4 ugP/L Summary The calibration curve is very linear over the range of standards. The replicates of sample #1 lie within the range of the standards, and their relative percent difference from each other is <15% (note also that they only differ by ~ 1 ugP/L, which is usually below the level of detection. This is excellent replication. The spike recovery for a 50 ugP/L spike was 95%, again very good, and well within the criterion of %. Conclusion – the data are valid! Conclusion: The data are valid % RPD = 100* (1.4)/ 11.9 = 12% % R = 100* ( )/50 = 95%

29 Total suspended solids and turbidity
Sediment plume off the south shore of Lake Superior Photo from NRRI

30 Total suspended solids and turbidity
TSS and turbidity are two common measures of the concentration of suspended particles. Suspended materials influence: Water transparency Color Overall health of the lake ecosystem Nutrient and contaminant transport Remember that neither method distinguishes between organic (plankton, detritus) and inorganic (e.g. clay) particles. However, the TSS filter may be ashed (combusted) at 475 to 550 oC and re-weighed to provide an estimate of the inorganic residue. The weight of the material that is vaporized represents the organic matter (OM) in the sample. It is also called: AFDW or AFDM – ash-free dry weight or mass TVS – total volatile solids VSS – volatile suspended solids POM – particulate organic matter Stream ecologists further fractionate organic solids into CPOM and FPOM (coarse and fine POM) because they are closely linked to the feeding ecology of benthic invertebrates.

31 Total suspended solids - sampling
TSS sampling in lakes involves collecting whole water samples No special handing or preservation is required but samples should be kept cool until analysis Recommended holding time is 7 days if kept at 4oC (but the sooner the better) Photo courtesy of NRRI

32 Total suspended solids - method
Filter a known amount of water through a pre-washed, pre-dried (at oC), pre-weighed (~ mg) filter Rinse, dry and reweigh to calculate TSS in mg/L (ppm)  Save filters for other analyses such as volatile suspended solids (VSS) that estimate organic matter Total Suspended Solids  TSS, or total suspended solids or total suspended sediment is pretty simple in theory. But, like most water quality parameters, it has some methodological problems and choices that require you to think before performing the measurement. The measurement is simple - separate the solids from the water using a piece of filter paper (actually you use a filter that looks like paper but is really made of glass fibers, pressed together; sort of like the fibers in the glass wool that is used for a fish tank). Then the filter plus the material on top of it are dried and weighed. If you remembered to weigh the filter before you used it (called “measuring the tare” or “taring” the filter), you can subtract this weight with the remainder being the weight of the solids. Some technique notes: 1. Filtration apparatus: A variety are available and vary considerably between the limnology/academic research community and the wastewater lab community. Fritted glass filter support bases work very well and provide a very uniform layer of material, but in time may clog. Our lab prefers plastic frits with fine slots, especially if you can get those with magnetic bases that allow you to place the filtration funnel on without a clamp. 2. You need to have enough particulate material on the filter to allow you to get a significant weight change (the more the better!), BUT if you try to filter too much water it could take you into tomorrow to filter it and the remember that the pore size is changing. Standard Methods (APHA) recommends that the final weight on the filter pad be 10 to 200 mg. Actual procedure notes: set the dried, weighed filter onto a filtering flask – wrinkled side up. shake the sample bottle to distribute particulate material uniformly and pour into a graduated cylinder (usually mL depending on TSS levels) APHA 1998 recommends that the final weight of the filter pad be 10 to 200 mg. pour into filtration funnel and turn on pump. Remember- the amount water to filter depends on TSS concentration so pour a bit at a time. rinse cylinder when empty with DIW and record volume - add more if the water is still filtering freely. filter until it slows to a trickle and you can see a definite brown or tan “spot” on the filter. turn off pump and bleed vacuum; carefully remove the filter and set it on a piece of paper towel to air dry if you just lay a wet filter on a smooth flat surface it may stick when it dries and in peeling it off to weight it you will lose some fibers and create an error. Laying them on a square inch of paper towel to air dry avoids this problem. Safest to put them in a drawer to keep them clean and protected from a sudden gust of air from a door opening. dry the filter at 103 to 105o C, let it cool to room temperature, and weigh it. Dry it, cool it, and weigh it again. Continue until the fiber reaches a constant weight. Record the end weight. With experience you can reduce this effort as long as you are consistent and follow an established protocol that is documented in your lab’s quality assurance/ quality control (QA/QC) manual.

33 Total suspended solids - method
What type of filter to use? It turns out that the filter you choose and how much water you filter may be important. Different brand filters have somewhat different nominal pore sizes and in fact these change as water is filtered and particles gradually plug up the pores. Unfortunately, this is not stated in standard reference manuals such as the various water quality methods “bibles” published by EPA, USGS, the American Public Health Association (see for list).    Under the Federal NPDES program, EPA method and Standard Methods 2540D are approved for measuring TSS in effluents or natural waters subject to regulatory requirements for discharges. Commonly used glass fiber filters include Whatman GF/F, GF/C and GF/A (from finer to coarser pores) Reeve-Angel 984-AH and 934 AH (finer-coarser) Gelman AE (approximately similar to Whatman GF/C’s) 4. Because of difference you may get in your TSS numbers just because of the filter you use (the NRRI Lab has found that values can vary by >50%), it is probably better to use the same filters year after year so that long-term trends will not be affected by methodology changes (“glitches” is the technical term). 5. Washing the filter is important because loose material can introduce an error. He filters must be pre-washed, dried and weighed to a constant weight (<0.5 mg variation or <4% of initial weighing). For non-regulatory routine monitoring work, do some preliminary checks to see how big these variations are to allow you to streamline your technique. After the washed filters are dried and tared, we store them in small plastic petri dishes (XX cms diam) that can be labeled with the filter # and weight and date and sample ID and even re-used later.

34 Total suspended solids
Some examples of filter types: Membrane filters retain sub-micron particulates and organisms Glass microfiber filters are made from 100% borosilicate glass Polycarbonate - offers precise pore size but reduced flow From Depth Filtration vs. Surface Membrane Filtration A depth filter traps contaminants both within the thickness of the filter and on the surface of the filter. Cellulose filter papers and glass fibers are depth filters with no defined pores, unlike a membrane that is a screen filter with a defined pore structure. Cellulose filter papers and glass fibers consist of a matrix of intertwined fibers in which particulates may be trapped in as well as on the surface of the filter. Since a depth filter does not have defined pores it is characterized not in terms of pore size, but in "particle retention". The advantage of a depth filter is that since entrapment and adsorption occur within the upper fraction of the media, there is a considerable surface area available for filtration. Also, because of its random matrix of fibers filter paper retains a large percentage of particulate. The disadvantages of a depth filter are that in the case of a sudden surge of differential pressure, as can occur when a vacuum is suddenly applied, the filter media will slough off fibres or particles during the filtration period. Also particulates trapped within the matrix can be forced through the matrix and contaminate the filtrate. In many applications a depth filter is used as a prefilter to clean a sample. A good example is the Whatman glass fiber filter paper GF/B with a thickness of 675 um and mean pore size of 1.3µm. A screen membrane traps particulates larger than its rated pore size on its surface. Particulates smaller than the specified pore size may either pass through the membrane or may be captured within the membrane. The advantages of a screen membrane are that rigid particulates can form a porous cake on the surface of the screen membrane and effectively improve the throughput of the screen. Also there is little risk of the filtrate being contaminated. The major disadvantage of a screen membrane is that the filtration process is slow compared with a depth filter Membrane filters are used for critical applications such as sterilizing and final filtration. An example is the PVDF membrane with 0.2µm pores size. A "combination filter" combines different membrane pore sizes or combines depth media and membrane filters to create a serial filtration units. These multi-layer filters can offer an economical alternative to using individual prefilters and final filters. A double filter membrane can have unique characteristics not achieved by either of the constituent single membranes. The filter membrane must be capable of retaining liquids during the application. Hydrophobic filters may be appropriate. Examples of such situations are: Cell capture prior to assay Removal of bacterial bio-load Retention of precious liquids

35 Total suspended solids – method
There are many different set-ups attach funnels by clamp, screw-on, or magnetic base plasticware useful in the field multiple towers Total Suspended Solids  Notes at by Tim Loftus       TSS, or total suspended solids, seems like an easy and innocent enough test to perform. For the most part, it is. But are you following this test procedure as it should be followed? Would the way you perform this test hold up in the legal system? You may get “workable” results for plant operations, but what about reporting these results to a regulatory authority? If you do not follow approved methods – exactly – then the results, even if they are accurate, may not be legally acceptable. These are not hypothetical situations. I have been through it, and fortunately, my results were not only accurate, they were performed legally. There was no argument.       Under the Federal NPDES program, you have two approved methods for TSS: EPA method and Standard Methods 2540D. It is important that you read the methods for the details of the test. But in general, both methods require that you wash, dry and weigh the glass fiber filters until you achieve a constant weight. A constant weight shows a variation of less than 0.5 mg or <4% of the previous weighing. Only then should you use the filter pads for TSS analysis. Believe it or not, a lot of junk gets washed off the filter pads. If it is not washed, a positive interference often results. The last thing you need when measuring TSS on your final effluent is a few extra milligrams that shouldn’t be there. It can make the difference between meeting your discharge limits or failing them. Wash and dry the pads for all your NPDES reporting and any other situations where the results are legally binding.       Sample size is equally important. Say, for instance, that you are running 50 ml of final effluent through the filter in the TSS test. To report the results in mg/l you must factor the measured TSS mg up by 20 times. In doing so, you are also factoring any error by at least 20 times. This can be significant if you have a low TSS limit in your NPDES permit. In this case, it would be better to use a much larger sample volume.       However, too much sample with high amounts of TSS can be harmful. A water-entrapping crust can form on the filter pad and will give a positive interference. This can happen especially with WAS and RAS samples. While these are not considered “legal” samples, the results do affect plant operation decisions. Standard Methods recommends that the final weight on the filter pad be 10 to 200 mg.       Another important aspect of TSS analysis is the filtration apparatus. EPA method requires that all filtration apparatus be fitted with a coarse (40-60um) fritted disk. Standard Methods requires only a filter apparatus with a reservoir to have a fritted disk. The fritted disk is a porous glass disk used to support the filter paper. It helps to provide equal suction under the whole filter pad. Many of the filtration funnels out there (Gooch, Buchner, membrane filter funnels) contain perforated disks or bottoms as filter supports. Realistically, perforated disks typically will perform well, but for many samples, they will not.       Finally, don’t forget to repeat the drying and cooling of the filter and sample until a constant weight is achieved.       These TSS method requirements mentioned are often overlooked in many places. However, it is important to do them. Read the EPA or Standard Methods test procedures over for the details. Performing the analysis correctly may take a little more time, but by doing so, your results will be accurate and they will be legal. And that is why we do lab work. Otherwise, why bother doing it?

36 Total suspended solids
Necessary TSS equipment Analytical balance Drying oven Filter and petri dish Other necessary equipment. Analytical balance must be accurate to g. Drying oven must be able to maintain temperature 103 to 105 oC. Images from NRRI

37 Total suspended solids
Calculate TSS by using the equation below: TSS (mg/L) = ([A-B]*1000)/C where A = Final dried weight of the filter (in milligrams = mg) B = Initial weight of the filter (in milligrams = mg) C = Volume of water filtered (in Liters)

38 How do turbidity and TSS relate?
A general rule of thumb: 1 mg TSS/L ~ NTU’s of turbidity BUT – Turbidity scattering depends on particle size so this is only a rough approximation How does turbidity relate to TSS ? Just some general information from lakes . Review-http://wow.nrri.umn.edu/wow/under/parameters/turbidity.html Also remember that plankton contributes to turbidity as well and that living cells are > 70 % water. A sample with high turbidity due to plankton may NOT correlate well to TSS as one with particulates due to erosional or resuspended silt. See also for a discussion focusing on stream turbidity and TSS.

39 Turbidity - meters Most use nephelometric optics and read in NTUs (nephelometric turbidity units) Field turbidity measurements are made with: Turbidimeters (for discrete samples) Submersible turbidity sensors (Note: USGS currently considers this a qualitative method) Laboratory instruments: Turbidimeters (bench models) References: National Field Manual for the Collection of Water-Quality Data. Techniques of Water-Resources Investigations Book 9:Handbooks for Water-Resources Investigations

40 Turbidity Turbidimeters Nephelometric optics
nephelometric turbidity is estimated by using the scattering effect suspended particles have on light detector is at 90o from the light source Reference: (from (Aug 2002)) Turbidity is due to: Inorganic particles such as clays and silts Organic material, both living and detrital, autocthonous and allocthonous, particulate and dissolved (e.g. dissolved color) The Relationship between Among JTU'S, and NTU's The Secchi depth measurement has been roughly correlated with Jackson Turbidity Units (JTU's). These units were based upon a standard suspension of 1000 parts per million diatomaceous earth in water. By diluting this suspension, a series of standards was produced. Jackson Turbidity Units (JTU's) are the application of these standards to the original device for measuring turbidity called the “Jackson tube.” The Jackson tube is a long glass tube suspended over a lit candle. A sample of water was slowly poured into the tube until the candle flame as viewed from above could no longer be seen. This device is no longer used because it is not sensitive to very low turbidities. A turbidimeter measures turbidity as nephelometric turbidity units (NTU). Instruments such as the turbidimeter that measure the scattering of light are called nephelometers. Both NTU's and JTU s are interchangeable units. They differ only in that their name reflects the device used to measure turbidity. However, it is now known that different optical geometries can produce somewhat different results and so this equivelance is really only an approximation.

41 Turbidity – units and reporting
Nephelometric Turbidity Units (NTU) Standards are formazin or other certified material JTU’s are from an “older” technology in which a candle flame was viewed through a tube of water 1 NTU = 1 JTU (Jackson Turbidity Unit) Photo from NRRI

42 Turbidity – formazin standards
Example of a set of formazin standards

43 Turbidity - Here is a range of NTUs using clay

44 Turbidity – meters and probes
Bench and portable instruments and kits vs. Submersible Turbidimeters YSI wiping turbidity YSI 6820 with unwiped turbidity Credits: (Note- these images are simply to illustrate the variety of laboratory turbidimeters that exist and is not meant to be a recommendation or endorsement of any particular manufacturer. However, the units used for most images in the WOW curriculum have been used by WOW staff in the field and/or laboratory. Bench an portable interments – sample is collected at a discrete depth - collect water sample and analyze water in Lab: (Left) Turner Designs Aguaflor Turbidimeter/Fluorometer (www.turnerdesigns.com ); (Right) Hach Kit and Bench Model (www.hach.com) Hydrolab

45 Turbidity - methods Comparability of different methods:
With the proliferation of automated in situ turbidity sensors there is concern about the comparability of measurements taken using very different optical geometries, light sources and light sensors. The US Geological Survey and US Environmental Protection Agency are currently (August 2002) developing testing procedures for a field comparison of a number of instruments produced by different manufacturers. We will include the USGS/EPA results when they become available. Standard Methods refers to : APHA Standard methods for the examination of water and wastewater. American Public Health Association, Washington, D.C.

46 Turbidity - calibration
Turbidity free water = zero (0 NTU) standard USGS recommends filtering either sample water or deionized water through a 0.2 um or smaller filter to remove particles WOW uses deionized water that is degassed by sparging (bubbling) with helium, to minimize air bubbles that may give false turbidity readings Notes: 1. Many turbidimeters use a 1 zero, 1 standard value calibration procedure. Set the zero using filtered lab DI water as a “blank”, then insert and read a selected calibration standard and adjust the instrument to agree with the “certified” standard. 2. WOW follows this procedure using a high end standard but prepares at least 1 additional standard near mid-range. We then use a linear regression routine to generate a standard curve from these 3-4 values for calculating sample values as per the standard curves for colorimetric nutrient analyses.

47 Turbidity - standards Standards range depends on anticipated sample values Lakes - typically 0-20 NTU Streams and wetlands , 0-50 or NTU 2 non-zero standards typically adequate (response is linear) Types of standards Formazin particles (either from a “recipe” or purchase a certified, concentrated stock solution -usually 4000 NTU) Other commercially available materials, e.g., polystyrene Formazin - need to worry about storage limits. Primary stock of 400 NTU’s lasts < 1 month when refrigerated. Dilute working standards from intermediate stock solution daily. See next slide.

48 Suggested holding times
Turbidity – standards Prepare daily 2 to 20 NTU Hach Company Prepare weekly All dilutions EPA Region 5 Standard Methods (APHA 1995) Prepare monthly 20 to 40 NTU Suggested holding times Concentrations Source Table of standards Needs references REFERENCES- 1. Hach Chemical Company,Loveland, CO, USA. 2. Standard methods = APHA Standard methods for the examination of water and wastewater. American Public Health Association, Washington, D.C. 3. EPA Region 5 = LIT CITE

49 Biochemical Oxygen Demand (BOD)

50 BOD BOD measures the amount of oxygen consumed by microorganisms as they decompose organic matter, as well as the chemical oxidation of inorganic matter The BOD test measures the amount of oxygen consumed during a specified period of time (usually 5 days at 20o C) References: EPA Stream Monitoring Manual: USGS Field Manual:

51 BOD 5 DO is measured initially and again after a 5-day incubation at 20o C BOD is computed from the difference between initial and final DO The rate of oxygen consumption is affected by a number of variables: temperature pH the presence of certain kinds of microorganisms the type of organic and inorganic material in the water

52 BOD – sample collection
Grab samples in clean, sterile containers (usually only surface sampling) If analysis is begun within 2 hours of collection, cold storage is unnecessary If analysis will be delayed > 24 hrs, store at or below 4o C Warm chilled samples to 20o C before analysis Note: Samples are usually only collected from the surface and in the nearshore zone, as well as in feeder streams, ditches, and outfalls. The assay is designed for high-strength wastewaters and so is not a routine lake monitoring parameter because of its lack of sensitivity. Appropriate parameters for estimating organic matter in unpolluted natural waters are: Dissolved organic matter (DOM) Particulate organic matter (VSS, POM) Dissolved and particulate carbon (DOC and TOC)

53 BOD - analysis Equipment needed: Incubation bottles
Air incubator or water bath thermostatically controlled at 20 +/- 1o C DO meter and probe Bottle preparation details are in BOD (PDF).

54 BOD Reagents: Dilution water – provides nutrients necessary for microorganism growth Seed – a population of microorganisms capable of oxidizing the organic matter in the sample Commercially available or freeze-dried culture A “conditioned” bacteria source (effluent from a biological treatment source such as a wastewater treatment plant). Glucose-glutamic acid standard See BOD (PDF) for details The dilution water consists of phosphate buffered deionized (or milli-Q) water that contains nutrients and minerals necessary to sustain the growth of microorganisms while they chow down on organic matter. In nature these would be supplied in the water. Phosphate buffer Magnesium sulfate Calcium chloride Ferric chloride Ammonium chloride Sodium sulfite Glucose-glutamic acid standard: Because BOD is essentially a bioassay, results can be influenced greatly by the presence of toxicants that are bacteriacidal or the use of poor seeding material. Periodically check dilution water quality, seed effectiveness, and technique by making BOD determination on pure organic compounds and samples with known additions. Use a mixture of 150 mg/L glucose and 150 mg/L glutamic acid as a standard check solution.

55 BOD – QA/QC Assure quality with:
Seed control – determine the BOD of the seeding source Dilution water blank – used to check for quality of unseeded dilution water and incubation bottle cleanliness Steps to Include: Read and record temperature of incubator Prepare replicate bottles for dilution water blanks and seed controls Include at least one set of replicate samples per analysis Residual DO- you do not want all of the DO to be used up in the samples because this will affect bacterial metabolism (the “seed”). Coming up with the right amount of dilution is almost an art. It helps to have had some prior experience with the waters you are analyzing. Seed control: note that the addition of the seed itself adds organic material to the sample. Reading the DO uptake in dilution water + seed and subtracting this BOD from the run will account for this.

56 BOD - procedure Blanks Prepare dilution water, bring to 20o C and aerate Add sufficient seeding material to produce a DO uptake of 0.05 to 0.1 mg/L in 5 d (dilution water) Samples Add sample to bottle and dilute. Dilutions should result in a residual DO of at least 1 mg/L and DO uptake of at least 2 mg/L after 5 day incubation Aerate dilution water by shaking the partially filled container or by aerating with organic-free filtered air. Protect quality by using clean glassware, tubing, and bottles. Samples: you made need to perform a range of sample dilutions to reach the desired DO uptake. Sample dilutions are made in the BOD bottle. Using a graduated cylinder. Add the desired sample volume to the individual BOD bottles of known capacity. Fill bottles with enough dilution water so that when you insert the stopper all of the air will be displaced leaving NO air bubbles. A good rule of thumb to use regarding dilutions is to have an overall depletion rate of ~ 1.0 mg O2/L/day. For an initial DO of 7 – 8 mg/L, this would yield a final value on Day 5 of about 2 – 3 mg O2 /L. This is a big enough change to measure accurately, but high enough to keep your bugs healthy and happy.

57 BOD – procedure Steps in procedure:
Fill bottles with enough dilution water so the stopper displaces all of the air, leaving NO air bubbles Read initial DO Incubate for 5 days at 20o C Read final DO Calculate BOD5 correcting for the exact duration Rinse DO probe between determinations to prevent cross-contamination of samples. The DO probe used for BOD analysis is specifically made to fit the opening in BOD bottles. It also has a vibrating mechanism that mixes the sample while reading. A gentle stirring with a stir bar may be used if necessary to move water past the probe’s membrane. Too vigorous mixing may introduce atmospheric O2 into your O2-depleted sample which produces an error in your determination.

58 BOD Calculations When dilution water is not seeded:
When dilution water is seeded: Where: D1 = DO of diluted sample immediately after preparation (mg/L) D2 = DO of diluted sample after 5 d incubation P = decimal volumetric fraction of sample used B1 = DO of seed control before incubation B2 = DO of seed control after incubation f = ratio of seed in diluted sample to seed in seed control = (% seed in diluted sample) (% seed in seed control)

59 Phytoplankton/Algae – counting methods
Photo from NRRI - This image was taken with a Nikon CoolPix 800 digital camera. We just held the lens up against the microscope eyepiece.

60 Algae- counting methods
Wet mounts Filter Counting chambers Utermohl requires an inverted microscope (light from above) Sedgewick rafter chamber Hemocytometer 3 main methods; Direct counts from raw water sample-only works for extremely high densities since the volume put on the slide is only ~ 0.05 to 0.1 mL Concentrate a relatively large sample ( mls) onto a fine membrane filter (cellulose acetate, cellulose nitrate, or mixed cellulose esters type of filters, usually 0.45 um pore size) using a filtering apparatus. A Utermohl settling chamber also concentrates cells by allowing them to settle (at least 24 hrs) down onto a glass coverslip. For waters with very low cell counts it may be necessary to settle 1-2 L sample down to about 100 mLs or so, then settling this concentrated volume in the Utermohl chamber.

61 Algae – counting methods
Microscopes capable of magnifications of 100X to 1000X Compound microscope Inverted microscope Less expensive inverted microscope

62 Algae- taxonomy Use an algal taxonomic key that shows species from your geographical area Phytoplankton are continually being described and re-classified so it’s essential for a good taxonomist to keep current (not easy by any means) It’s a good idea to take photographs of slides for cataloging Links: REFERENCE KEYS: For a list of nationwide references and on-line images go to - Smithsonian Environmental Research Center Hard copy: Prescott, G.W How to know the freshwater algae, 3rd ed. Wm. C. Brown Co, Dubuque, IA. Prescott, G. W Algae of the western Great Lakes Area, revised ed. Otto Koeltz Science Publishers, W. Germany. Dillard, G.E Common Freshwater algae of the United States. An illustrated key to the genera (excluding diatoms). 173 p, 298 figs, 21x16cm, spiral bound. ISBN $35.00 Phycological Society of America has lots of links (http://www.psaalgae.org/) On-Line: Susquehanna University Algal Image Archive at Cedar Eden Environmental, LLC at Great Lakes Diatoms Microscopy UK Images of freshwater algae from the Keewenaw Peninsula, Michigan at Introduction to chrysophytes at

63 Algae – determining biomass
Algal biomass (standing crop): A quantitative estimate of the total mass of living organisms within a given area or volume Biovolume estimates: Identification to genus and species level Calculate cell volume by approximation to nearest geometrical shape Count cells over a known area of the slide so cells per unit volume can be determined Chlorophyll

64 Algae – determining biovolume
Taxonomic keys often include questions about size Determining size is basically like using a ruler. The standard ruler for a microscope is called an "ocular micrometer," which is fitted into the eyepiece of your microscope If you don’t have a micrometer, you can use the diameter of the field. Each of these methods requires that you first standardize your microscope against a ruler of known length; at low magnification, this standard could be a transparent office ruler, but at higher magnifications a stage micrometer is needed. Be aware that different microscopes are not exactly the same and the size goes down with increased magnification (i.e.: the more you magnify something, the smaller the actual item is compared to the image you see).

65 Algae – determining biovolume
Some formulas to estimate biovolume from cell dimensions (Wetzel & Likens 2000) Rod B A Sphere Ellipsoid Wetzel, R. G. and G.E. Likens Limnological Analysis, 3rd Ed. Springer-Verlag New York, NY, Inc.

66 Algae – chlorophyll determination

67 Algae – chlorophyll determination
Measuring chlorophyll-a concentration remains the most common method for estimating algal biomass Chlorophyll-a concentration has also been shown generally, when comparing lakes, to relate to primary productivity (Wetzel 1983) Can be used to assess the physiological health of algae by examining its degradation product, phaeophytin The degradation product phaeophytin has been shown to contribute 16-60% of the chlorophyll-a content in seawater and freshwater (APHA 1998). However, in practice, it is often difficult to make sense from routine phaeophytin data. More sophisticated pigment and fluorescence techniques have also been developed for estimating algal abundance, community structure, growth, and physiological status.

68 Algae – chlorophyll basics
Algal biomass is most commonly estimated by chlorophyll-a. Units are ug/L or mg/L (ppb and ppm) Detection limit depends upon method used An in-depth microscopic enumeration of the dozens of species of algae present in a water column each time a lake is sampled is prohibitively costly and technically impossible for most monitoring programs. Further, in many lakes a large portion of the algal biomass may be unidentifiable by most experts (these are appropriately called LRGTs or LRBGTs -- little round green things and little round blue-green things). However, measuring the concentration of chlorophyll-a is much easier and provides a reasonable estimate of algal biomass. Chlorophyll-a is the green pigment that is responsible for a plant's ability to convert sunlight into the chemical energy needed to fix CO2 into carbohydrates. To measure chlorophyll-a, a volume of water from a particular depth is filtered through a fine glass-fiber filter to collect all of the particulate material greater than about 1 micron (1/1000th of a millimeter) in size. The chlorophyll-a in this material is then extracted with a solvent (acetone or alcohol) and quantified using a spectrophotometer or a fluorometer.

69 Algae – chlorophyll methodology
Spectrophotometry and fluorometry, utilizing 90% acetone extraction, remain the most commonly used methods Spectrophotometry is most widely used but fluorometry is more sensitive and may be used when low levels of chlorophyll are anticipated or when handling large volumes of water is logistically difficult

70 Algae – chlorophyll sampling
0 to 2 m integrated samples are usually collected for chlorophyll analysis Samples must be kept cool and out of direct sunlight until filtered Freeze moist filters until analysis Note: There is a school of thought that filters can or should be dried prior to freezer storage. However, this remains a minority opinion and is not recommended in standard agency lab manuals and guidelines.

71 Algae – chlorophyll instrumentation
Spectrophotometer: Visible with 1-2 nm bandwidth Matched cuvettes, 1-5 cm Fluorometer: Requires excitation and emission filters specifically for chlorophyll measurement Photos from NRRI

72 Algae – chlorophyll filtration
Apparatus - extraction  Prewashed 47 mm glass fiber filters (GF/C, GF/F, AE, or equivalent) Gelman polycarbonate filtration tower or equivalent Vacuum pump (5 to 7.5 psi) Centrifuge (clinical) DIW/acetone (90%) washed 15 mL Corex centrifuge tubes with caps Notes: Lots of other filtration set-ups are available. We (NRRI) like the Gelman magnetic base with no clamp. There are many other extraction solvents including: ethanol (ETOH), methanol (METH), and combinations of these with dimethyl sulfoxide (DMSO). The extrations may also be hot or cold. We are presenting 90% acetone here because it remains the most commonly used method for US state monitoring programs.

73 Algae – chlorophyll filtration (cont.)
Filter a known volume of water through a GF/C filter Volume filtered depends upon algal density Add a few drops of saturated MgCO3 solution near the end When all the water has been pulled through, fold the filter into quarters and wrap in foil We generally filter enough water to get good color on the filter (green or brown or somewhere in between). For some lakes only a few hundred milliliters will do but for an ultra-oligotrophic lake like Lake Superior you may need to filter 2 liters or more! Three things degrade chlorophyll a : Light Heat Acid So, keep the sample out of direct sunlight, keep it cool (store frozen), and do not touch the filter with your fingers. Your fingers are slightly acidic so use a forceps to handle the filter. The saturated MgCO3 buffers the sample.

74 Algae – chlorophyll storage
Wrap the folded filter in a square of foil, label, then freeze Record the volume filtered, date, site, depth, replicate # all with permanent marker Store the filter in the freezer at < 20o C EPA holding time for a frozen chlorophyll filter is 2 weeks

75 Algae – chlorophyll extraction & analysis
Tear filter into several pieces Place in a test tube Add 10 mLs of 90% acetone Extract overnight at 4oC Chlorophyll analysis: After hr extraction, centrifuge to settle filter debris Read absorbance or fluorescence of the supernatant The glass fiber filters, if left suspended in the extract, will contribute to increased turbidity in the sample which can reduce the chlorophyll signal. To clarify the sample, centrifuge for 20 minutes at 4000 rpm. Cap the tubes to prevent evaporation of the acetone during the extraction and analysis. Details in chlorophyll method (PDF). Sometimes periphyton samples contain blue-green or green algal filaments. Often the chlorophyll within these filaments will not extract completely without physical disruption. This can be done using a tissue grinder but care must be taken not to overheat the sample (remember that heat degrades chlorophyll). Place the filter in the grinder, then place the grinder into a beaker filled with crushed ice. An alternative is to use an ultrasonic disrupter or sonicator. Alternative solvents and solvent mixtures have been shown to be better for some groups of algae (such as hot methanol for blue-greens). However, 90% acetone extracted overnight at 4o C or ground and analyzed after a 0.5 to 1.0 hr extraction are considered “standard methods”.

76 Algae – chlorophyll measurement
Measure absorbance of a 90% acetone solution blank at 750 nm and at 664 nm to correct for primary pigment absorbance Record sample absorbance at 750 nm and 664 nm Estimate phaeophytin by acidifying the sample. Record the absorbance at 665 nm and again at 750 nm Run working standard solutions of purified chlorophyll-a (Sigma Chemical Co. Anacystis nidulans by the procedure used for the blank) Chlorophyll-a and phaeophytin concentrations in the extract are determined spectrophotometrically using the standard method for chlorophyll-a in the presence of phaeophytin (as detailed in Standard Methods 1992). A Perkin-Elmer Lambda 3B UV/VIS spectrophotometer with a 1 nm spectral bandwidth and optically matched 4 cm micro-cuvettes are used at NRRI for routine analysis. To measure phaeophytin, acidify the 4 mL of solution in the cuvette to a final molarity of 3 x 10-3 by adding 100 ul of 0.12N HCl to the 4 mL of extract in the cuvette. Allow acid to react for 60 seconds and record the absorbance at 665 nm and again at 750 nm. This is a correction made for the 665 nm wavelength (for phaeophytin). Using purified Anacystis chlorophyll is convenient and allows a known concentration to be made. However, it should still be “calibrated” using standard equations and using a calibrated spectrophotometer. Alternatively, spinach or lettuce or even grass, can be extracted overnight in 90% acetone to produce a solution of largely chlorophyll that can be measured and used as a check standard or calibration standard for fluorometry.

77 Algae – chlorophyll and phaeophytin
What is phaeophytin? Degradation product of chlorophyll Absorbance wavelength (665 nm) is very close to that of chlorophyll (664 nm) H acid The addition of acid to a chlorophyll sample replaces the Mg atom from the center of the molecule with a hydrogen atom to form phaeophytin. This degradation product of chlorophyll can interfere with the determination of chlorophyll because it absorbs light and fluoresces in the same region of the spectrum. There are actually several related degradation products and if present in substantial amounts significant errors in chlorophyll a values will occur. So, if the algal community was healthy and growing happily when sampled, there will be less phaeophytin in the sample. When acid is added to the extract, there should be a significant drop in absorbance indicating that most of the pigment present initially was chlorophyll a which was then converted to phaeophytin by the acid. If the algal community was senescing or growing old, the change in absorbance after the acid addition will be smaller indicating that there were more degradation products present in the sample.

78 Algae –spectrophotometry calculations
Where: b = before acidification a = after acidification E664b - [{Abs664b(sample)–Abs664b(blank)}-{Abs750b(sample)–Abs750b(blank)}] E665a - [{A665a(sample)-Abs665a(blank)}-{Abs750a(sample)-Abs750a(blank)}] Vext = Volume of 90% Acetone used in the extraction (mL) Vsample = Volume of water filtered (L) L = Cuvette path length (cm)

79 Algae – chlorophyll QA Quality assurance
There are no commercial QA check standards Lab replicates are usually not done Essentially, the analysis is a one-shot deal, you don’t get a second chance, so be careful Field replicates should be done every 10 samples Cut filters in half and save one half if nervous Patchiness is a term often used in aquatic ecology. In essence it means that you cannot count on organisms and chemical constituents to be evenly distributed in space or time within a lake. In very productive lakes with surface scums, field replicates can be quite different. Larger sample volumes and integrating samplers can help reduce this variability.

80 Periphyton Photo for section change

81 Periphyton Collection:
Qualitative grabs or scrapings versus quantitative sampling from a known surface area Different methods are used for collecting periphyton from rocks, woody debris, macrophytes, bottom substrates or other substrates

82 Periphyton – in situ sampling
Resulting material from a rock scrub (to the right) containing: Macro invertebrates Detritus Fungi Bacteria as well as algae More on how to actually determine periphyton biomass is found in Modules 4/5 and 9.

83 Periphyton – sample prep
Here’s a portion of the previous sample after being deposited on a glass fiber filter in preparation for chlorophyll extraction or AFDW determination. This involves filtering a known volume of the periphyton suspension. Remember that this volume is arbitrary and what really matters is the weight that was collected from a known area of substrate.

84 Periphyton – biomass estimation
Wet weight Dry weight (dried at 103–105o C) Ash free dry weight (AFDW) Loss on ignition (LOI) Combust at o C Chlorophyll (extract as per phytoplankton) Particulate organic carbon and/or nitrogen (POC or PON) Muffle furnace Remember that this measurement does not distinguish between detritus, algae, fungi, bugs or bacteria. Methods closely follow those used for water column algae. Note that for the organic matter estimation, it is the material that is burned off that is the organic matter. The residue is inorganic “ash”.

85 Periphyton – biomass calculations
Once you have a measure of chlorophyll or AFDW you’ll need to calculate per unit area. Note: You have to be careful to record the various volumes of solvents and dilution factors and what % of the original sample was used for the different analyses. For water samples it’s easy – just note the volume analyzed. For periphyton and sediment, there was a total amount of sample collected for a given surface area. A portion, by weight, was used for each analysis.

86 Periphyton biovolume Measure cell dimensions with an ocular or stage micrometer to calculate cell volume. Sphere A Ellipsoid B Rod Wetzel, R. G. and G.E. Likens Limnological Analysis, 3rd Ed. Springer-Verlag New York, NY, Inc. This is really difficult for some periphyton communities that have complex mixtures of different algal groups. On the positive side, sometimes the sample may be dominated by a few species such as filamentous “glops” that are seen in quiescent waters.

87 Bacteria – E. coli and fecal coliforms
REFERENCES 1. US Geological Survey (USGS), Water Resources--Office of Water Quality, National Field Manual Chapter 7.1 FECAL INDICATOR BACTERIA 2.

88 Bacteria – E. coli and fecal coliforms
Fecal bacteria are used as indicators of possible sewage contamination These bacteria indicate the possible presence of disease-causing bacteria, viruses, and protozoans that also live in human and animal digestive systems E. coli is currently replacing the fecal coliform assay in most beach monitoring programs These indicator bacteria are generally not considered harmful by themselves although some can be pathogenic (disease causing).

89 Bacteria - indicators The most commonly-tested fecal bacteria indicators are: total coliforms fecal coliforms Escherichia coli (E. coli) fecal streptococci and enterococci All but E. coli include several species of bacteria E. coli is a single species in the fecal coliform group Fecal coliforms, a subset of total coliform bacteria, are more fecal-specific in origin. E. coli is a species of fecal coliform bacteria that is specific to fecal material from humans and other warm-blooded animals. It has many strains however, some of which may be very toxic to humans. EPA currently recommends E. coli as the best indicator of health risk from water contact in recreational waters; some states have changed their water quality standards and are monitoring accordingly. Others are in the process of analyzing for both E. coli and fecal coliforms concurrently before switching to E. coli in order to generate conversion factors.

90 Bacteria – EPA standards
The U.S. EPA recommended standard for E. coli concentration in recreational waters: The geometric mean for > 5 samples within a 30-day period shall not >126 E. coli colonies per 100 ml of water; and No sample > 235 E. coli colonies/100 ml of water in a single sample For fecal coliforms: Geometric mean for > 5 samples within a 30-day period not > 200 cfu/100mL < 10 % of samples > 400 cfu/100 mL in any 30-day period Notes: EPA’s BeachNet program is an excellent source of information on this topic (http://www.epa.gov/waterscience/beaches/) as is the Center for Disease Control (CDC) at For Minnesotans, check out the new Lake Superior Beach Monitoring Program at

91 Bacteria – 2 indicator methods
Two basic methods: membrane filtration 2. multiple-tube fermentation Images taken from: 1.) Document Title: Petri Dish Category:Health-Related Research and Technologies, Biomolecular Systems and Science, Date of Image/ Photo: August 7, 1982 Background: URL of this page: 2.) Membrane filtration involves filtering several different-sized portions of the sample using filters with a standard diameter and pore size, placing each filter on a selective nutrient medium in a petri plate, incubating the plates at a specified temperature for a specified time period, and then counting the colonies that have grown on the filter. This method varies for different bacteria types (variations might include the nutrient medium type, the number and types of incubations, etc.). The multiple-tube fermentation method involves adding specified quantities of the sample to tubes containing a nutrient broth, incubating the tubes at a specified temperature for a specified time period, and then looking for the development of gas and/or turbidity that the bacteria produce. The presence or absence of gas in each tube is used to calculate an index known as the Most Probable Number (MPN). EPA’s On-Line Methods for Microbiology Detailed procedures for analyzing samples for bacterial, viral, and protozoan pathogens.

92 Bacteria – membrane filter technique
The fecal coliform MF procedure uses an enriched lactose medium and incubation temperature of 44.5 ± 0.2o C for selectivity. Results in 93% accuracy (APHA 1995) in differentiating between coliforms found in the feces of warm-blooded animals and those from other environmental sources. Fecal Coliform is reported as colony forming units per 100 mL (CFU/100 mL). Standard Methods sections in microbiological examinations to be read prior to performing this procedure include: Quality Assurance, Laboratory Apparatus, Washing and Sterilization, Samples (Collection, Storage and Preservation) and Membrane Filter Technique for Members of the Coliform Group.

93 Bacteria – membrane filter equipment
Materials needed for MF method: Air incubator or water bath Non-corrugated forceps Heat sterilizer (Bacti-Cinerator) Filter flask and tower (Autoclavable) Vacuum pump or water aspirator Incubator or water bath needs to be thermostatically controlled at 44.5 ± 0.2o C.

94 Bacteria – membrane filter equipment
MF materials (continued): Sterile 50 mm petri plates (with tight-fitting lids) Sterile 0.45 um gridded membrane filters Sterile absorbent pads Autoclave (121o C at psi) Images from: Incubator or water bath needs to be thermostatically controlled at 44.5 ± 0.2o C.

95 Bacteria – membrane filter procedure
Saturate the absorbent pad with M-FC broth Select a sample volume that will yield colonies/filter Filter sample and dilution water through pad Place pad into petri dish Invert plates and place in incubator for 24 hrs Procedure From: Ameel, J. et al Analytical chemistry and quality assurance procedures for natural waters, wastewater, and sediment samples. NRRI/TR-98/28 Place a sterile absorbent pad in a sterile petri dish and pipet mL of the M-FC Broth to saturate the pad. Pour off any excess liquid before the membrane filter is placed in the plate. Select a sample volume which will yield colonies/filter. When the bacterial density is unknown filter several volumes to ensure a countable density of colonies. Use a sterile filtration unit at the start of each filtration series to avoid contamination. Using sterile forceps, place a membrane filter (grid side up) on the porous plate of the filter tower. Place the funnel over this and lock into place. Filter a minimum volume of 20 mL. When less than 20 mL of sample is to be filtered, add approximately 10 mL sterile dilution water to the funnel before filtration then pipet the sample into the funnel and filter the entire dilution. This will aid the distribution of the colonies over the entire filter. Rinse the interior of the funnel with approximately 10 mL buffered dilution water three times. Immediately remove membrane filter with sterile forceps, and place it in a petri dish using a rolling technique to avoid air bubbles under the filter. Plates must be incubated within 30 minutes of filtration. Invert the plates and place in a water tight bag in a water bath at 44.5 ± 0.2o C. If Klebsiella strains are present the temperature may be raised to 45o C. A “pre” and “post” blank (sterility check) must be analyzed with each run. If more than 10 samples are to be analyzed, a sterility check is run after every 10 samples.

96 Bacteria – membrane filter counting
Fecal coliform colonies bacteria are various shades of blue. Non-fecal colonies are gray to cream colored. normally, few of these are present. See examples from

97 Bacteria – MF counting (cont.)
image showing method of counting

98 Bacteria – multiple tube fermentation
MTF image process E coli and fecal coliform counts in Lakes Washington and Sammamish are available on the King County lakes website.

99 Bacteria – cleaning and sterilizing
All equipment Wash equipment thoroughly with dilute nonphosphate, laboratory-grade detergent. Rinse 3 X with hot tap water Rinse again 3-5 X with deionized or glass-distilled water. Glass, polypropylene, or Teflon™ bottles If sample will contain residual chlorine or other halogens, add Na2S2O3. If sample will contain > 10 ug/L trace elements, add EDTA. Autoclave at 121 C for 15 min or bake glass jars at 170 C for 2 hrs. Stainless-steel field units Flame sterilize with methanol (Millipore™ Hydrosol units only), or autoclave, or bake at 170 C for 2 hrs Portable submersible pumps and pump tubing Autoclavable equipment (preferred): autoclave at 121 C for 15 min. Non-autoclavable equipment: Submerge sampling system in a 200 mg/L laundry bleah solution and circulate solution through pump and tubing for 30 min; follow with thorough rinsing, inside and out, with sample water pumped from the well. **SEE NOTES **DO NOT USE THIS METHOD TO DISINFECT EQUIPMENT USED TO COLLECT SAMPLES FOR SUSEQUENT DETERMINATIONS OF TRACE ELEMENTS AND ORGANIC SUBSTANCES.

100 Bacteria – USGS summary
Test (media type) Ideal count range (colonies per filter) Typical colony color, size, and morphology Total coliform bacteria (m-Endo) 20-80 Colonies are round, raised and smooth; 1 to 4 mm di; and red with golden-green metallic sheen. Escherichia coli After primary culture as total coliform colonies on m-Endo (NA-MUG) None given but much fewer in number than total coliforms on the same filter Colonies are cultured on m-Endo media as total coliform colonies. After incubation on NA-MUG, colonies have blue florescent margins with a dark center. Count under a long wave ultra violet lamp in a completely dark room. Fecal coliform bacteria (m-TEC) 20-60 Colonies are round, raised and smooth with even to lobate margins; 1 to 6 mm di; light to dark blue in whole or part. Some may have brown or cream colored centers. Escherichia coli (m-TEC) Colonies are round, raised and smooth; 1 to 4 mm di; yellow to yellow brown; many have darker raised centers. Fecal streptococci (KF media) 20-100 Colonies are small, raised, and spherical; about 0.5 to 3 mm di; glossy pink or red in color. Enterococci (m-E and EIA) Colonies are round, smooth and raised; 1 to 6 mm di; pink to red with a black or red dish – brown precipitate on underside.

101 Fecal coliforms – troubleshooting
Uneven; not mixed well; low volume poor seal around the edges; poorly seated with air bubble Dry spot from poor seating No matter which assay is used, after incubation there should be ~20-60 colonies evenly distributed across the Petri dish

102 Fecal coliforms – troubleshooting (cont.)
Too many – use less sample Too few – use more sample Looks good

103 Aquatic vegetation Aquatic vegetation sampling is detailed in Module 8 Part C Lake sampling References

104 Aquatic vegetation – biomass method
Harvested material is sorted by species Stripped of periphyton Weighed, dried at o C and reweighed Biomass is usually expressed as wet weight or dry weight per m2 Dried material may be ground and subsampled for organic matter, %C, %N, %P or other constituents

105 Aquatic vegetation – biomass method
A separate set of carefully pressed and dried specimens may be set aside for archives A blotted, but wet subsample may be extracted for chlorophyll. The wet:dry ratio is important for comparing areal chlorophyll values to other parameters

106 Zooplankton KEYS AND OTHER REFERENCES
Dodson, S.I., and D.G. Frey Cladoceran and other Branchiopoda. Pp in Thorp, J.H., and A.P. Covich (eds.). Ecology and classification of North American freshwater invertebrates. Academic Press, San Diego. Pennak, R Freshwater invertebrates of the United States. Wiley, 3rd edition. Williamson, C.E Copepoda. Pp in Thorp, J.H., and A.P. Covich (eds.). Ecology and classification of North American freshwater invertebrates. Academic Press, San Diego. Balcer, M.D., N.L. Korda, and S.I. Dodson Zooplankton of the Great Lakes: a guide to the identification and ecology of the common crustacean species. University of Wisconsin Press, Madison. 48 pp. Stemberger, R.S A guide to rotifers of the Laurentian Great Lakes. U.S. Environmnetal Protection Agency, Cincinnati, Ohio. (Available from National Technical Information Service, Springfield, Virginia. PB ) A work in progress: The smallest cite on the web

107 Zooplankton – sample preservation
Most commonly 95% ethanol or 5% formaldehyde (formalin) Animals preserved in formalin sometimes become distorted which complicates size measurements. One solution involves the addition of 40 g/L sucrose to the 5% formaldehyde. Rose Benegal dye is also used by many to stain the critters for ease of identification Use formalin with care in a ventilated area or under a hood = suspected carcinogen and a confirmed irritant to lungs and eyes. The sucrose solution is very messy and sticky.

108 Zooplankton – equipment
1. Hensen Stemple pipettes Compound microscope 6. Sedgwick-Rafter counting slide 2. Dissecting microscope 5. Folsom Plankton Splitter 3. Ward Counting Wheel 4. Hensen-Stemple pipettes - used to subsample a full sample. Usually the volume of the sample is mLs which is too much volume and way too many animals to count. Note: you can easily reduce the volume of water in a sample by wrapping a small piece of zoopankton netting around a cutoff pasteur pipette or an automatic pipette with a rubber band For slide chambers that hold a specified volume of sample be sure to mix the full sample well before sub-sampling a small aliquot. Best to count several aliquots and average them. Splitter and wheel – minimizes the “settling problem” when sub-sampling. Dissecting scope for “coarse identification” Compound scope for examining leg hairs, mouth parts and other details needed to distinguish taxa at the species level or for measuring body length for biomass estimation. If the sample doesn’t have more than a few hundred animals it is most accurate to count everything to avoid errors and the need for replicating aliquots associated with sub-sampling. All B/W images from WildCo.com

109 Zooplankton – taxonomy
Taxonomy is complex so ID to species level is best left to the experts but genus and order level are relatively easy As with phytoplankton, organism size is important to determine Photo from -

110 Zooplankton – detailed biomass
Daphnia pulex Approximate sizes (not to scale) 2 mm 0.5 mm Cyclops 1 mm Typically need animal densities (#’s /liter) and body lengths grouped by species, genus, or order. Then you need an equation relating body length to length from the literature if it is not possible to do this yourself.

111 Zooplankton –total biomass
Total community biomass may be estimated by simply measuring the wet weight (or dry weight) of the zoops from a given tow with known volume. Leptadora

112 Zooplankton – biomass example
To determine # animals/L you need to determine the volume of water filtered through the net. Example Using a Wisconsin net with a small, 13 cm diameter opening for a 0 to 5 m vertical tow: Where d = 0.13 m and z = 5.0 m = 0.66 m3 = 66 liters

113 Benthic samples Photo from NRRI

114 Benthic samples Processing benthic invertebrate samples
Determining sediment bulk characteristics: Texture (% sand, silt, clay) % organic matter Total carbon, nitrogen, and phosphorus concentration Sediment oxygen demand

115 Benthic invertebrates – sample processing
Sorting into taxonomic groups, Identifying to desired taxonomic level, Data entry Image from:

116 Benthic invertebrates – sample processing
Rinse the sample in a 500 m mesh sieve to remove and fine sediment. Sticks and leaves can be visually inspected and then discarded. 1 cm opening will hold back larger material like leaves, sticks, shells, but still allow organisms through. Wash the sample through with water Check the large material, the stuff that eventually gets discarded, for any attached or burrowed organisms Wash the material that passes through the sieves into a jar. This is best done in the field when sampling sediment.

117 Benthic invertebrates - sub sampling
Spread the sample evenly across a pan marked with grids Randomly select 4 squares, remove the material and preserve in jars Sub-sampling is not a requirement and is actually frowned upon by some scientists but it reduces the effort required for sorting and identification. You are, of course, assuming that the organisms are spread equally across the grid. This may or may not be the case. This technique does not work well for species occurring in very low numbers. The sub-sample is sorted and preserved separately from the remaining original sample for quality control checks. Any counts made must be multiplied by the sub-sample fraction to calculate densities (in this example X 4).

118 Benthic invertebrates – identification
Most organisms are identified to the lowest possible taxonomic level Lowest taxonomic level depends on the goals of the analysis, expertise, and available funds Taxonomic references: There are very many but here are some classics: Meritt, R.W. and K.W. Cummins (editors) An introduction to the aquatic insects of North America, 3rd ed. Kendall/Hunt Publishing Company, Dubuque, Iowa. Pennak, R.W Freshwater invertebrates of the United States, 3rd ed. J. Wiley & Sons, New York. See pgs 7-20 through 7-33 in: Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling Rapid Bioassessment Protocols for Use in Streams and Wadeable Rivers: Periphyton, Benthic Macroinvertebrates, and Fish, 2nd Edition. EPA 841-B s. U.S.

119 Benthic invertebrates – data processing
Metric An attribute with empirical change in value along a gradient of human influence In other words, a measurement made to determine if humans have had an impact in a natural system. Index An integrative expression of site conditions across multiple metrics. An index of biological integrity is often composed of at least 7 metrics. (Karr and Chu 1997) Karr, J. R. and E.W. Chu Biological Monitoring and Assessment: Using multimetric indexes effectively. EPA 235-R

120 Benthic invertebrates - data metrics
Many metrics have been developed for aquatic invertebrates. Richness measures Composition measures Tolerance measures Much more detail can be found in: Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling Rapid Bioassessment Protocols for Use in Streams and Wadeable Rivers: Periphyton, Benthic Macroinvertebrates, and Fish, 2nd Edition. EPA 841-B s. U.S. See Chapter 9, Biological Data Analysis. Trophic/habitat measures

121 Benthic sediment – bulk properties

122 Sediment - bulk properties
Texture % organic matter Total carbon Organic matter Nutrient content: Bioavailable phosphorus Total phosphorus Total nitrogen Sediment oxygen demand

123 Sediment - texture Refers to the shape, size, and three-dimensional arrangement of the particles that make up sediment Gravels and pebbles can be measured using calipers Sand is measured using sieves of different mesh size Silts and clays are more difficult Silts and clays: The most common method is called elutriation where the sample is treated with a dispersant within a cylinder. The finest fractions stay in suspension longest, whereas the heavier fractions sink more rapidly. Samples are withdrawn at a given depth at specific intervals, then dried and weighed. Direct measurement of clay particles is only possible using an electron microscope. (adapted from: 9/23/03) Of course there are much more quantitative methods (this is a well developed science). For much more detail see the following: USGS USGS EAST-COAST SEDIMENT ANALYSIS: PROCEDURES, DATABASE, AND GEOREFERENCED DISPLAYS

124 Sediment - % organic matter
Measured as mg/g sediment % carbon may also be important to measure, particularly in studies of sediments contaminated with pesticides, PAHs, and dioxide Methods closely follow those used for water column algae. Note that for the organic matter estimation, it is the material that is burned off that is the organic matter. The residue is inorganic “ash”.

125 Sediment – phosphorus content
Potentially bioavailable P from sediment or sediment traped material can be estimated from a single extraction with 0.1 N NaOH. Total P can be extracted using persulfate or hot HCl acid procedure. Both procedures involve extracting P into a solution which is then analyzed for P content using the ortho-P ascorbic acid method. Bioavailable P method : NRRI Method IV.3.20 Total P for sediment : NRRI Method IV.3.22 Note: There are many more sophisticated P – fractionation from sediment protocols in the peer-reviewed literature. These are simply a few methods that have proven useful in the NRRI lab over the past 15 years.

126 Sediment – C:N content Coming soon

127 Sediment – exchangeable NH4+
Coming soon

128 Sediment – oxygen demand
Coming soon

129 18 4 4


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