Freshwater Ecology
Laboratory Manual
Kansas State University
Division of Biology
BIOL 612
Fall 2014
Professor: Walter Dodds
Teaching assistants: Matt Trentman, Allison Veach
Lab manual prepared by: MJ Bernot, WK Dodds, R Bernot, JM O’Brien, A. Riley, JN Murdock, D.
Larson, M Trentman, and A Veach
Table of Contents
Page Number
1.
Lab Schedule .............................................................................................................................................. 3
2.
Lab Requirements ....................................................................................................................................... 4
3.
Lab Notebook Checklist……………………………………………………………………………………5
4.
Individual Lab Protocols
a) Lab 1 Introduction, Beginning Field Equipment, and Kimball Pond
Introduction………………………………………………………………………………… 6
Equipment Introduction ........................................................................................................ 6
Dissolved Oxygen ................................................................................................................. 7
b) Lab 2 Tuttle Creek Reservoir……………………………………………………………………... 9
c) Lab 3 Pottawatomie State Fishing Lake II………………………………………………………. 9
d) Lab 4 Beginning organism ID and use of microscopes
Organism Identification ........................................................................................................ 10
Planktonic Organisms……………………………………………………………………10
Microscope Introduction ................................................................................................. 10
Zooplankton Enumeration ..................................................................................................... 12
e) Lab 5 Lake models and properties of water
Lake Models.......................................................................................................................... 13
Properties of Water ............................................................................................................... 14
f) Lab 6 Stream invertebrates – McDowell Creek & Kings Creek
Aquatic Insect Order Distinguishing Characteristics………………………………………. 15
g) Lab 7 Alkalinity, P-I curve
Alkalinity .............................................................................................................................. 16
Photosynthesis-Irradiance Curve .......................................................................................... 18
h) Lab 8 Island Biogeography .............................................................................................................. 19
i) Lab 9 Introduction to the Spectrophotometer and Colorimetric Analyses, SRP, Chlorophyll a
SRP…………………………………………………………………………………………. 22
Chlorophyll a………………………………………………………………………………..23
j) Lab 10 Autoanalyzer, NO3-………………………………………………………………………… 25
k) Lab 11 Water and Sewage Treatment...………………………………………………………….. 27
l) References………………………………………………………………………………………….....28
2
Lab Schedule
Day
Date
Subject
Tues
26 Aug
Lab 1. Equipment introduction; Trip to Kimball Pond; Winkler, meet behind fire
station at Claflin and Kimball at 1:30, be prepared to get muddy
Tues.
2 Sep
Lab 4. Organism identification; Microscope introduction
Sat.
6 Sep
Lab 2, 3. Pott State Lake II and Tuttle Creek Reservoir 8:30 - ~2:00 (or camping @
Pott II 8:00 pm Fri), meet in front of Bushnell, be prepared to get wet, need drivers
with State Park permits
Tues.
9 Sep
Lab 5. Lake models, hydrodynamics, organism identification (work on Lab 1)
Tues.
16Sep
Lab 6. Field trip, McDowell Creek, Kings Creek, meet in front of Bushnell, be
prepared to wade, need drivers Identify Stream Organisms
Tues.
23 Sep
Free week
Tues
30 Sept
Lab 6 (cont). Identify Stream Organisms
Tues.
7 Oct
Lab 7. Alkalinity; Winkler, P-I
Tues.
14 Oct
Lab 8. Pillsbury Crossing, island biogeography, meet in front of Bushnell, be prepared
to wade, need drivers. first notebook grading, see points for required elements!
Tues.
21 Oct
Lab 8 (cont). Count stream organisms
Tues.
28 Oct
Lab 9. Intro to Spectrophotometer, SRP, chlorophyll, NH4+ assay
Tues.
4 Nov
Lab 10. NH4+ assay using an auto-analyzer
Tues.
11 Nov
Lab 11. Tour sewage treatment and water treatment, meet in front of Bushnell, need
drivers
Tues.
18 Nov
Final nutrient analyses and organism ID
Tues.
25 Nov
Thanksgiving break
Tues.
2 Dec
Discuss field results; Lab cleanup; Practical review; finalize lab notebooks, final
analyses
Tues.
9 Dec
Notebooks due, practical exam
3
Laboratory Requirements:
Many professional ecologists, fisheries biologists, and environmental scientists are required to keep a field/laboratory
notebook as part of their expert duties. This is an actual record of events, not something to be constructed after the fact,
although information can be added (such as analysis). Lab books are professional records of your work; they are
considered legal records and have been subpoenaed in courts of law. Thus, we hope to build good professional scientific
work habits with this course requirement.
The purpose of the lab notebook is to record data and field observations as well as to document results and state
conclusions. You will not be graded on neatness, but results must be readable. Take as many field notes as you can
because they frequently come in handy when analyzing data upon return to the lab. Although there are specific
requirements for the lab notebook, it only needs to be organized and neat to the extent that the T.A. can grade it and you
can refer back to it and interpret what it says (the table of contents will aid in finding information).
The laboratory/field notebook is required for completion of the course (i.e. part of your lab grade). General rules to
follow:
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Notebooks should have bound pages (not spirals, not loose leaf binders)
Each notebook should have a table of contents (leave the first few pages blank for this)
All pages should be numbered and dated
For each lab, indicate what is to be accomplished (i.e. objectives, methods, etc.)
After collection of data written in lab notebook, data should be transferred electronically to a Microsoft Excel
spreadsheet and appropriate graphs generated. As hard copies can sometimes get lost, scientists always back up data
electronically!
Your notes should be written in ink at the time of the lab or field trip. Do not copy over notes after lab or field
trip. Notes written in ink can serve as legal documents. Note: ethanol can cause ink to run and smear. We highly
recommend backing up the important field and lab notes on the computer in case the notebook gets lost or damaged.
Note you can scan at library to pdf format relatively easily.
Be sure to compare methods of sampling or analysis where relevant.
In every lab that requires graphs to be created (most labs), graphs should be generated in Microsoft Excel (or other
graphical software). Details on how this should be done will be explained in class. When submitting lab notebooks
to be graded, also include a binder or folder with printed graphs (include axis labels and units when necessary).
In EVERY lab in which organisms are observed: sketch organisms and show indication of scale. At least 15 species
of algae and 20 species of invertebrates (zooplankton, insects, protozoa, and crustacea) should be sketched during
the semester.
Grading requirements are listed in this lab manual.
4
* 1st lab notebook check due
Freshwater Ecology Lab Book
Grade
Points
Possible
Points
Freshwater Ecology Lab Book
Grade
Points
Possible
Points
Objectives
20
______
McDowell Creek and Konza
Results & Conclusions
Ammonium/SRP/Nitrate+Nitrite
Alkalinity
Discharge
Methods
20
______
2
2
2
______
______
______
Kimball Pond -Results/Graphs
Dissolved Oxygen*
Temperature*
Light*
PH*
1
1
1
1
______
______
______
______
Water & Sewage Trt Plants
Results & Conclusions
NH4+ , NO3, SRP
3
______
Notes
2
Conductivity*
1
______
Ammonium
SRP
Nitrate+Nitrite
Chlorophyll a
Alkalinity*
Conclusions
Pott II State Lake -Results/Graphs
Dissolved Oxygen*
1
1
1
1
1
10
______
______
______
______
______
______
1
______
Temperature*
Light*
PH*
Conductivity*
1
1
1
1
______
______
______
______
Ammonium
SRP
Nitrate+Nitrite
Chlorophyll a
Alkalinity*
Conclusions
Tuttle Creek Reservoir
Results/Graphs
Dissolved Oxygen*
Temperature*
Light*
PH*
Conductivity*
1
1
1
1
1
10
______
______
______
______
______
______
1
1
1
1
1
______
______
______
______
______
Ammonium
SRP
Nitrate+Nitrite
Chlorophyll a
Alkalinity*
Conclusions
1
1
1
1
1
10
______
______
______
______
______
______
Organism I.D
Algae & Macrophytes
number
Invertebrates
15
______
Number
20
______
Plankton Count
(including plots with depth)
P-I Curve
Standard Curves
Ammonium
SRP
2
______
2
______
5
5
______
______
Pillsbury Crossing
Island Biogeography
20
______
Lake Models
Results & Conclusions*
10
______
Site Comparison
10
______
Total
200
______
5
Lab 1. Introduction and Beginning Field Equipment
A considerable amount of technical skill is required to be a successful aquatic scientist. You must be able to
characterize biotic and abiotic aspects of aquatic environments. The abiotic characteristics of water bodies can provide a
great deal of insight into the factors affecting the system. Factors such as light, water chemistry, O2 availability,
temperature, and pH can have extreme effects on the biotic community. These factors can also indicate anthropogenic
effects on the system. Water, because of its dynamic nature, is especially vulnerable to pollution inputs. Such inputs can
alter the nutrients available in the system or in the physical environment. The physical environment can sometimes
result in drastic effects on the surrounding biotic community. Water pollution can cause eutrophication,
methelhemoglobemia, and toxic algal blooms.
We will be sampling three lakes and three streams for several limnological parameters. The sampling sites are
Kimball Pond, Tuttle Creek Reservoir (Lab 2), Pottawatomie State Fishing Lake II (Lab 3), Kings Creek (Lab 6),
McDowell Creek (Lab 6) and Pillsbury Creek (Lab 8). We will characterize these sites according to abiotic and biotic
parameters to enable us to understand and compare the dynamics of each system.
A student who successfully completes the laboratory for this class should be able to: 1) produce a reliable
field/laboratory notebook, 2) utilize basic limnological equipment, 3) understand the principles behind basic chemical
analyses of aquatic systems, 4) identify some of the most common organisms found in lakes and streams, and 5)
integrate field data with theory learned in class lectures.
Objectives, Lab 1:
1. Become familiar with the protocols for use of standard limnological equipment and oxygen titration.
2. Compare 2 methods of DO determination (Winkler Method vs. DO meter)
3. Measure limnological parameters in Kimball Pond.
4. Describe the biology of a hypereutrophic farm pond.
Requirements, Lab 1 (note, some of these will be done in later labs).
1. Notes on use of standard equipment, learn to use equipment.
2. Show calculations for at least one Winkler oxygen analysis and plot oxygen concentration against pond depth.
Compare Winkler method to DO meter methods.
3. Plots of temperature, conductivity, pH, O2, light, light attenuation coefficients, redox, alkalinity, ammonium,
nitrate, SRP, and chlorophyll a against depth.
4. List of organisms seen at each site.
Equipment
Secchi Disc
 weighted disc, 20 cm in diameter with alternating white & black pies
 used to measure transparency or water clarity
 black marks = 10 cm, or 0.1 m (units: meters)
Li - Cor Photometer
 radiation sensor connected to meter
 used to measure light, either as radiant flux (quantity of electromagnetic energy over time, quanta s-1 watts s-1, or
joules s-1) or as irradiance (radiant flux per unit area, µ mol quanta m-2 s-1 or µ Einst m-2 s-1)
 will need to calculate the Light Attenuation Coefficient (LAC, or η) at depth x ( in meters below surface):
LACx = ln (light at surface) - ln (light at x)
depth x
Van Dorn Bottle
 open ended cylinder that shuts on contact with messenger
 used to sample water chemistry and phytoplankton at different depths
Eckman Dredge
 square box with spring-operated jaws that shut on contact with messenger
 used to sample soft sediments and invertebrates
 marked every 1 m
Schindler-Patalas Sampler
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
clear, plastic boxed trap allowing quantitative sampling of water and plankton at specific depths
pH Meter
 used to measure the negative of the log of the molar concentrations of H+ in solution: pH= -log[H+]
 values can range from acidic (0-5) to alkaline (9-14)
 most unpolluted waters have a range from 6-9
Redox Meter
• redox (Eh), or oxidation-reduction potential (ORP) is the number of free electrons
Conductivity Meter
 estimates total dissolved solids (TDS) held in solution or the resistance of a solution to electrical flow
 TDS = (0.6 x alk.) + Na + K + Ca + Mg + Cl + SO4 + SiO3 + (NO3-N) + F
 units: mho cm-1 or Siemens cm-1
 freshwater range 50-1500 µmho cm-1, DI water 0.5-4.0 µmho cm-1
 units displayed by the meter are mmho cm-1
Plankton Tows
 fine mesh nets; mesh opening of the plankton nets range from 1050-67 µm depending on the net used, most are
about 200 µm for phytoplankton and 400 µm for zooplankton
 used to sample zooplankton
 can be used vertically or horizontally through water column
Dissolved Oxygen (DO) Meter
 measures DO
 calibrate for temperature, salinity, and altitude
Alkalinity, Ammonium, Nitrate, SRP, and Chlorophyll a
• See later labs
Dissolved Oxygen
"Oxygen is the most fundamental parameter of lakes, aside from water itself" -Wetzel, 1983
Why is dissolved oxygen (DO, O2) so important when it occurs in such low concentrations in water relative to air (a few
mg L-1)? It determines biological and biochemical reaction (redox) that occurs in the environment and it is essential to
metabolic functions of aerobic organisms. DO affects the solubility of inorganic nutrients and determines in what
direction reactions take place (redox potential).
Where can oxygen dissolved in lakes originate? Dominant sources are the atmosphere and photosynthesis. Respiration
and chemical reactions consume O2.
What environmental factors influence DO? Temperature, barometric pressure, and concentration of ions.
Methods of determination:
1. Bunsen: boil out gases at atmospheric pressure or lowered pressure; most accurate, but difficult and not plausible
for field measurements.
2. Colorimetric methods: most are inaccurate.
3. DO meters: more expensive; must be calibrated carefully for accurate results, but easy to use.
4. Winkler: can be more accurate and reliable than DO meters, but is more cumbersome.
Winkler Method
Procedure (see detailed protocol for more information):
1. Fill 300 mL BOD bottle without bubbles (3x the volume, filling the bottle is an art, you need to be able to do this
without introducing air bubbles, make certain you learn this)
2. Add 2 mL Mn SO4 below the surface
3. Add 2 mL alkaline-iodide-azide reagent (NaOH + KI) below the surface
-oxygen binds to iodide
4. Stop bottle and invert several times
5. When 1/2 of the precipitate is settled, add 2 mL concentrated H2SO4 below the surface
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6. Invert bottle several times
7. When precipitate is dissolved, transfer 200 mL to 500 mL beaker
-important to transfer exact volume
8. Titrate standard sodium thiosulfate solution with burette until pale straw color
9. Add a few drops of soluble starch solution
10. Mix solution to get a uniform blue color
11. Calculate approximate DO concentration:
1 mg O2 L-1 = 1 mL of titrant
Winkler Method: Detailed Protocol - Azide Modification
1. General Discussion
Under alkaline conditions, dissolved O2 in the sample will oxidize the Mn2+ to form MnO2, which will form a
brown precipitate. H2SO4 is then added to create acidic conditions, allowing the MnO2 to then oxidize the I- to I2 causing
the solution to become yellow-brown in color. Thiosulfate is then used to titrate the I2 back to I-, completion of this
reaction is indicated by the starch solution. Thus by measuring the amount of sodium thiosulfate needed to remove the I2
from the solution, we can calculate the amount of O2 originally in the solution. Sodium azide is used to prevent
interference of this reaction by NO2- and dissolved Fe. Use the azide modification for most wastewater effluent and
stream samples, especially if samples contain more than 50 µg NO2- - N/L and not more than 1 mg ferrous iron/L. Other
reducing or oxidizing materials should be absent. If 1 mL KF solution is added before the sample is acidified and there is
no delay in titration, the method is applicable in the presence of 100 to 200 mg ferric iron/L.
2. Reagents
a. Manganous sulfate solution: Dissolve 480 g MnSO4 •4H2O, 400 g MnSo4•2H2O, 364 g MnSO4•H2O in distilled
water, filter, and dilute to 1 L. The MnSO4 solution should not give a color with starch when added to an acidified
potassium iodide (KI) solution.
b. Alkali-iodide-azide reagent:
1) For saturated or less-than-saturated samples – Dissolve 500 g NaOH (or 700 g KOH) and 135 g NaI (or
150 g KI) in distilled water and dilute to 1 L. Add 10 g NaN3 dissolved in 40 mL distilled water.
Potassium and sodium salts may be used interchangeably. This reagent should not give a color with starch
solution when diluted and acidified.
2) For supersaturated samples – Dissolve 10 g NaN3 in 500 mL distilled water. Add 480 g sodium hydroxide
(NaOH) and 750 g sodium iodide (NaI), and stir until dissolved. There will be a white turbidity due to
sodium carbonate (Na2CO3), but this will do no harm. CAUTION—Do not acidify this solution because
toxic hydrazoic acid fumes may be produced.
c. Sulfuric acid, H2SO4, conc.: One milliliter is equivalent to about 3 mL alkali-iodide-azide reagent.
d. Starch: Use either aqueous solution or soluble starch powder mixtures.
To prepare and aqueous solution, dissolve 2 g laboratory-grade soluble starch and 0.2 g salicylic acid,
as a preservative, in 100 mL hot distilled water.
e. Standard sodium thiosulfate titrant: Dissolve 6.205 g Na2S2O3•5H2O in distilled water. Add 1.5 mL 6N NaOH or
0.4 g solid NaOH and dilute to 1000mL. Standardize with bi-iodate solution. Note, we will not do this step in
class.
f. Standard potassium bi-iodate solution, 0.0021M: Dissolve 812.4 mg KH(IO3)2 in distilled water and dilute to
1000mL.
g. Standardization – Dissolve approximately 2 g KI, free from iodate, in an Erlenmeyer flask with 100 to 150 mL
distilled water. Add 1 mL 6N H2SO4 or a few drops of conc H2SO4 and 20.00 mL standard bi-iodate solution.
Dilute to 200 mL and titrate liberated iodine with thiosulfant titrant, adding starch toward end of titration, when a
pale straw color is reached. When the solutions are of equal strength, 20.00 mL 0.025M Na2S2O3 should be
required. If not adjust the Na2S2O3 solution to 0.025M. Note, we will not do this step in class.
h.
Potassium fluoride solution: Dissolve 40 g KF•2H2O in distilled water and dilute to 100 mL.
3. Procedure
a. To the sample collected in a 300-mL bottle, add 2mL MnSO4 solution, followed by 2 mL alkali-iodide-azide
reagent. If pipettes are dipped into sample, rinse them before returning them to reagent bottles. Alternatively, hold
pipette tips just above liquid surface when adding reagents. Stopper carefully to exclude air bubbles and mix by
inverting bottle a few times. When precipitate has settled sufficiently (to approximately half the bottle volume) to
leave clear supernate above the manganese hydroxide floc, add 2.0 mL conc H2SO4. Restopper and mix by inverting
several times until dissolution is complete. Titrate a volume corresponding to 300 mL original sample after
8
correction for sample loss by displacement with reagents. Thus, for a total of 4 mL (2 mL each) of MnSO4 and
alkali-iodide-azide reagents in a 300-mL bottle, titrate 200 × 300/(300-4) = 202 mL.
b. Titrate with 0.025M Na2S2O3 solution to a pale straw color. Add a few drops of starch solution and continue
titration to first disappearance of blue color. If end point is overrun, back-titrate with 0.0021M bi-iodide solution
added dropwise, or by measured volume of treated sample. Disregard subsequent recolorations due to catalytic
effect of nitrite or to traces of ferric salts that have not been complexed with fluoride.
4. Calculation
a. For titration of 200 mL sample, 1mL 0.025M Na2S2O3 = 1 mg DO/L.
b. To express results as percent saturation at 101.3 kPa, use the solubility data in Table 4500-O:I (Standard Methods).
Equations for correcting solubilities to barometric pressures other than mean sea level and for various chlorinities
(salinity) are given on p. 4-101 in Standard Methods.
5. Precision and Bias
DO can be determined with a precision, expressed as a standard deviation, of about 20 µg/L in distilled water and about
60 µg/L in wastewater and secondary effluents. In the presence of appreciable interference, even with proper
modifications, the standard deviation may be as high as 100 µg/L. Still greater errors may occur in testing waters having
organic suspended solids or heavy pollution. Avoid errors due to carelessness in collecting samples, prolonging the
completion of test, or selecting an unsuitable modification.
Lab 2. Tuttle Creek Reservoir
Situated in Riley, Pottawatomie, and Marshall counties in northeast Kansas, Tuttle Creek Reservoir was
constructed and is operated by the U.S. Army Corps of Engineers. The reservoir covers about 12,350 acres, averaging
about 1 mile in width. The dam is located on the Big Blue River five miles north of Manhattan. Completed in 1962, the
purposes of the reservoir are flood control, recreation, fish and wildlife conservation, water quality control, and
navigation supplementation.
Objectives
1. Measure limnological parameters of Tuttle Creek Reservoir
2. Describe the biology of a large impoundment
Sampling Protocol
Use standard limnological equipment (the tools you learned how to use at Kimball pond) to characterize mid-lake and
near-shore habitats of a reservoir.
1. Mid reservoir profile - Collect a vertical profile (2-meter increments) of the abiotic and biotic parameters; including
surface measurements such as light attenuation, and bottom measurements such as benthic invertebrates.
2. Near shore sampling - Characterize the biotic components of the littoral area of a reservoir.
Requirements:
1. Field notes
2. See lab grading sheet for all requirements on this laboratory
Lab 3. Pottawatomie State Fishing Lake II / Wetland
Pottawatomie State Fishing Lake II (Pott II) is located 7miles NE of Manhattan, Kansas in Pottawatomie
County. The lake was created from 1953 to 1955 by the Kansas Department of Wildlife and Parks on land generously
donated by Dr. and Mrs. Robert L. Freidrich. It covers 75 acres with a small wetland area on the northeast side that
resulted from the creation of the lake. In 1959, two Bigfoot sightings occurred near the shores of Pott 2. In one account,
a man saw a “real hairy man” standing beside his house. He drove across lawn and ditch to escape. At about the same
time, a woman reported seeing a “wildman” run into trees beside a field where she was working (Chronological List of
Bigfoot Sightings: http://www.netcomuk.co.uk/~rfthomas/cb/1951.html).
Objectives
1. Measure limnological parameters of Pottawatomie State Fishing Lake and adjacent wetland.
2. Describe the biology of a small impoundment.
3. Describe the biology of a wetland.
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Sampling Protocol
Lake - Use standard limnological equipment (same procedure as in Tuttle Creek Reservoir) to characterize midlake and near-shore habitats.
Wetland - Sample a longitudinal transect from the lake into the wetland area (3-4 points). At each point collect a
vertical profile (surface, mid depth, and bottom) of O2, redox, and pH. If a mat of Chara is present, be sure to
collect a profile within it. Also at each point, collect one zooplankton sample (Schindler trap) and one benthic
invertebrate sample (Eckman dredge).
Requirements:
1. Field notes
2. See lab grading sheet for all requirements on this laboratory
Lab 4. Beginning organism identification, use of microscopes
Objectives
1. Identify and enumerate organisms found in various lakes and streams
2. Learn major parts of microscope
3. Learn proper microscope technique
4. Calibrate microscopes
Requirements:
1. Organisms we are likely to find are listed below, the final list of organisms that you are responsible for ID in
practicum will be compiled near the end of semester
2. Need to show work on scope calibration
3. Will be tested on microscope parts and operation
4. Need to count at least one zooplankton sample per person
Organism Identification
Objective:
Note, we will see these throughout the course, not just in the first organism lab. This list will be subject to change
before the practicum depending on what we actually find.
Why Study Planktonic Organisms?
Much of the plankton is made up of or is dependent on primary producers (phytoplankton). Autotrophic phytoplankton
use light as an energy source to reduce CO2 to organic carbon through photosynthesis. Much of the oxygen in the
atmosphere so necessary for human life on earth was produced by primary producers. Zooplankton are heterotrophs that
depend on primary producers for food. Together, they constitute a large portion of the bottom two trophic levels in any
aquatic food web. We will become familiar with 5 groups of phytoplankton and several groups of zooplanktonic animals.
Planktonic Organisms
Zooplankton:
Insecta:
Diptera (Chaoboridae & Chironomidae)
Crustacea: Cladocera (Daphnia & Bosmina)
Copepoda (Cyclopoda, Harpacticoida & Calanoida)
Phytoplankton/ Benthic Producers:
Prokaryota: Phylum Cyanophyta (Blue-Green Algae or Cyanobacteria, includes Anabaena,
Oscillatoria, Nostoc, and Synechoccus)
Eukaryonta: Phylum Chlorophyta (Green Algae, includes Desmids, Spirogyra, Cladophora, Chara,
and Volvox)
Phylum Chrysophyta (Yellow-Green Algae, includes diatoms)
10
Phylum Euglenophyta (Euglena)
Phylum Pyrrhophyta (Dinoflagellates, includes Ceratium)
Microscopes
Vernier Scale: used to mark the position of an object on a slide, so one can easily return to that object
Magnification = eye piece x objective (e.g. 10 x 40 = 400)
Proper Focusing Technique
1. Focus on cover slip edge
2. Adjust field diaphragm to make small spot
3. Sharpen spot with condenser focus
4. Center spot in field of vision with condenser lens control knob
5. Open field diaphragm
*make sure condenser is closed
Calibrating Ocular Micrometer to Measure Organism Size (document in your laboratory notebook)
Use ocular micrometer (marked glass that fits inside ocular) and stage micrometer
1. measure ocular µm with stage µm by lining up a line on the stage micrometer with a line on the ocular micrometer
(one stage micrometer division = 10 µm)
2. if there is not an ocular micrometer available, use field width to standardize
3. repeat for each magnification
4. use same scope each time after calibrated
5. INDICATE SIZE LENGTH AND WIDTH FOR ALL ORGANISMS IDENTIFIED
Example:
You need to determine the number of µm/ocular divisions and divide.
995 µm/95 ocular divisions = 10.5 µm/ocular division at 10X magnification
If an organism at 10X is 3 ocular divisions long it would be 31.5 µm long.
Plankton Enumeration
The plankton community of freshwater lakes varies significantly from year to year as well as from season to
season (Lampert & Sommer 1997). Furthermore, vertical structuring of the community is likely during periods of lake
stratification (Leibodand & Tessier 1991). Many factors such as resource quantity and quality, food web structure, and
physical parameters may ultimately influence dynamics. Therefore, long-term monitoring of plankton in conjunction
with other long-term biotic and abiotic monitoring protocols may shed light on the dominant mechanisms associated
with such a highly variable system.
11
The composite biomass of phytoplankton populations can be estimated by measuring the concentration of
chlorophyll a pigments in water samples
We will be enumerating zooplankton by using dissecting scopes and a 5 mL counting wheel. For each picture,
identify the organism as far taxonomically as possible (not size and length). The quantitative count will be performed
only on Cladocerans, Copepods, and the Nauplii (larval stage of Copepod). Count all the organisms using the procedure
below and report.
1. Bring sample to 100 mL.
2. If already >100 mL, let settle for 10 minutes and pipette off excess H2O.
3. Stir sample in beaker with volumetric pipette (this pipette holds 5 mL of sample) for 15 seconds (until well mixed).
4. Take subsample (5 mL) with pipette.
5. Pour subsample into groove in counting wheel.
6. Let sample settle into groove for a minute or two (note: large amounts of algae may require more settling time).
7. Use dissecting scope to count all Cladocera, Copepoda and copepod Nauplii in groove by slowly rotating wheel.
Report zooplankton as number per liter:
# of zoops L-1
=
C × V’
V’’ × V’’’
where:
C = # of organisms counted
V’ = volume of the concentrated sample (100 mL)
V’’ = volume counted (5 mL)
V’’’ = volume of water through which net was towed (L) = πr2d
(r = radius of net opening, d =depth)
Note: if Schindler Trap was used, the calculations will need to be modified.
Lab 5. Lake Models and Properties of Water
Objectives:
1. Use model for temperate lake from Wetzel and Likens (1991). Take data of temperature versus depth, and
describe why you observe the results you do.
2. Examine several properties of water through experimental observation.
Requirements:
1. Raw data on lake model experiment
2. Explain why there is an exponential decrease in temperature with depth with a single heating episode in a lake
(before first wind)
3. Explain why thermal stratification sets up (describe interaction between thermal heating and intermittent mixing
by wind)
4. Observations on demonstrations of properties of water
5. Compare and contrast molecular and turbulent diffusion
6. Describe the effect of streamlining on flow boundary layer
7. Explain why the corn syrup demonstrations simulate water movements at very small scales, and what this
means for very small organisms
12
Lake Models
“The hydrodynamics of water movement is an integral component of a functional lake system. The importance of water
movements and associated turbulence either has been underestimated or neglected in a large majority of limnological
investigations. Consideration must be given to the effects of water movements on stratification and on the distribution of
temperature, dissolved gases and nutrients, and biota (Wetzel & Likens)."
The laboratory exercises outlined below are based on a simple model of a lake; keep in mind that you are studying a
model and not an actual lake. Furthermore, many of the formulas are simplified accordingly. Some virtues of the models
are:
1. Size is reduced to manageable proportions.
2. Complexity is reduced to the extent that mechanisms and processes may be appreciated readily and intuitively
understood.
3. Experimentation is facilitated, whereas it may be difficult and/or expensive with a natural system.
On the other hand, unjustifiable extrapolations to the natural condition could be made from the model if the observer is
not careful to distinguish between the two. In the experiments outlined here, for example, if the sides and bottom of the
aquarium were not insulated, there could be more rapid heat exchange between the model “lake” and its surroundings
than ever would be the case in nature. Likewise, the amount of energy supplied by the heat lamp in relation to the depth
of water is much higher than in natural situations. The reason for the modification is, of course, to accelerate the
process. These and similar points should be stressed in your reports.
These exercises are based on and modified from material prepared by Mortimer (1951) and Vallentyne (1967).
Caution: Take care not to splash water on the heat lamp----otherwise, EXPLOSION!!!
Lake Model 1: Temperate Lake
Purpose:
Thermally induced density stratification commences in the spring and persists through the summer for moderately deep
lakes in the Temperate Zone. These conditions are illustrated in this model. It is instructive in this exercise to separate
the actions of sun and wind, and to that extent the model is unnatural. The exercise also illustrates the phenomena of
internal seiches and convective sinking during night or autumnal cooling.
Procedures:
1. Prepare a chart with columns for time (minutes elapsed) and temperatures at depths of 1, 3, 5, 9, 13, 17, and 21 cm.
Leave a space at the extreme right for inserting temperatures at other (variable) depths.
2. Fill the aquarium with cold tap water and mix to 4°C with ice. Allow the system to come to rest (3 to 5 min) with a
small amount of ice floating on the surface, then introduce a moderate wind (first wind) on the top, blowing from
one side and then the other. After a minute or two you will notice that the temperature at 1 cm has dropped to 2 to
3°C. Stop the wind and record the temperatures at the surface and at each fixed depth. Next, remove the excess ice
as gently as possible and turn on the heat lamp. Record this time as the beginning of the experiment. With the light
on and no wind, record temperatures after approximately 5 min and every 5 min thereafter for approximately 50
min, taking special care to get additional temperature readings in regions where temperatures change rapidly with
depth (e.g., upper 5 cm). Plot temperature changes as a function of depth on graph paper as they are recorded. You
will observe an exponential decrease in temperature with depth. Why?
3. With the lamp still on, drop some crystals of catechol violet randomly over the surface. Allow them sufficient time
to sink so that most of the descending trails are in the upper third of the aquarium with a few extending into the
middle third, and still fewer to the bottom. Observe the positions of the trails and the complexities of the water
motion, particularly near the surface. What can you infer about the water currents in the “lake”?
4. Now introduce a light to moderate wind (second wind) blowing first from one side and then the other so as to mix
the surface waters into one homothermal mass. Avoid too strong a wind, which could destroy the thermocline that
you are now creating. The epilimnion (with homogenous distribution of color) should now be about 5- to 7-cm
thick. Turn off the light and allow the currents to slack; record the depth of the homogeneous red layer and
temperatures as a function of depth.
5. Leave the light off, if cool, remove from working area. Using the same wind speed (third wind), impart a slope of 10
to 15° to the thermocline by blowing from one side only. This slope can be achieved best in stages, allowing
successive winds with intervening periods of quiescence to build up the internal seiche. When the desired slope to
the thermocline has been obtained, stop the wind and measure the periods of the internal seiche (average
13
approximately five measurements). The oscillating period of the internal seiche should be recorded directly,
simply observe and time the rocking of the stained layer. Allow the system to come to rest, record the
temperatures at each depth, and measure the depth of the homogeneous red layer.
6. Carefully add a layer of ice cubes to the surface with minimal disturbance of the water. It will be necessary to cover
about three-fourths of the surface if small cubes or crushed ice are used. Observe the descending convection cells
sinking through the thermocline region, which in turn cause the thermocline to descend. When the thermocline has
descended to about mid-depth in the tank, carefully remove the remaining ice with as little turbulence as possible.
With a wind (fourth wind) from one side, build up a 10 to 15° slope on the thermocline and redetermine the periods
of seiches. Record temperatures and the depth of the homogeneous red layer.
7. Allow the system to rest and then with a strong wind (DO NOT SPLASH WATER ONTO THE HEAT LAMP) turn
the lake over. When mixing is complete, record the temperatures.
Properties of Water
Chemical Diffusion
20 µL of dye will be added to several agar plate wells at different times before class begins. Observe the movement of
dye within the agar plate. Calculate diffusion rate of dye by using Fick’s Law (see lecture materials and text) and
concentration information from lab.
Flow Boundary, Streamlining
1. Get two pieces of modeling clay the same size (about 3cm3 each)
2. Make one into a cube and one into a fusiform (torpedo) shape
3. Get a recirculating chamber with a variable speed and put a blob of dyed water on the bottom of the chamber
4. Slowly turn up the velocity, observe the dye and record observations
5. Place the clay cube in the bottom of the chamber, get an intermediate flow going
6. Use a pipette to introduce dye into the chamber and indicate turbulent and laminar flow (draw in lab book)
7. Repeat with fusiform shape and draw observations
8. Attach a piece of Cladophora (if samples are available) to the bottom of a recirculating chamber. Slowly turn up the
water velocity. Describe what happens to the shape of the algal thallus. Draw what happens to the shape at 0 and
high water velocity.
Reynold's Number (Re) Experiment
Work in groups of 2 or 3. We will use two media: water and corn syrup. On the scales that bacteria operate, water has the
properties of corn syrup. This experiment will consist of several treatments repeated with both media.
1. Stain 10 mL of water and 10 mL of corn syrup with food coloring, use lots, make it dark
2. Fill a large beaker with water and another with corn syrup
3. Pull a test tube brush through the corn syrup, and another through the water, note the differences in the amount of
force required
4. Add 2 mL of corn syrup halfway between the center and the edge of the beaker, halfway between the top and the
bottom with a pipette
5. Carefully stir three complete circles, using a glass rod, keeping the rod exactly 1 inch from the edge of the beaker,
then exactly reverse the mixing without removing the rod
6. Repeat using water and dyed water
7. Note the different responses to mixing
8. Clean equipment
9. Fill a graduated cylinder with water
10. Roll a marble in the dyed water solution and drop into cylinder
11. Observe what happens to the dye trail
12. Repeat with corn syrup and a marble rolled in stained corn syrup
13. Observe what happens to the dye on the marble
14. Let sit for 1 hour and observe again
14
Lab 6. Stream Invertebrates – McDowell Creek & Kings Creek
Why Study Aquatic Insects?
Benthic macroinvertebrates dominate the biomass in the sediments of most freshwaters and represent an
important component of productivity (Downing 1984). They can be important components of the food web because
some can digest cellulose and lignin, which other organisms cannot. Insects are medically important (e.g. Diptera) both
because of their sometimes-harmful (and irritating) bites and the potential for being vectors of diseases. Many aquatic
insects are used as environmental indicators (e.g. Plecoptera). Lastly, aquatic insects are useful for recreational fishing as
bait and lures.
Less than 10% of all insects are aquatic because of the difficulty in obtaining oxygen in aquatic environments
and the need to maintain ionic balance. Despite this small percentage, thirteen orders of insects need water during at least
some part of their life cycle (see Appendix for detailed dichotomous key). We will become familiar with all the aquatic
insect orders and several genera and species through collection, identification, and drawing.
Objectives:
1. Identify non-insects to Class and insects to Order (both scientific and common names).
2. Draw organisms and use keys provided.
Stream Invertebrates
Aquatic Insect Order
Common name
Ephemeroptera
Mayflies
Plecoptera
Stoneflies
Trichoptera
Caddisflies
Diptera
True flies
Hemiptera
True bugs
Coleoptera
Beetles
Odonata
Dragon flies
Megaloptera
Dobbson flies/Fish flies
Lepidoptera
Moths
Neuroptera
Spongillaflies
Orthoptera
Grasshoppers
Hymenoptera
Bees/wasps
Collembola
Spring-tails
Distinguishing Characteristics
3 "tails" on abdomen; abdominal gills
2 "tails" on abdomen; no abdominal gills
no wing pads; abdomen ends in pair of anal claws
head capsule, chewing mouth parts, no wing pads
mouth is in the form of a "beak", or elongated structure
larvae do not have prolegs; adults have platelike covering
large prehensile labile
abdomen in single filament or hooked prolegs
fleshy leg-like structures w/ hooks on abdomen
long slender projection on mouth for sucking
enlarged hind legs (grasshopper)
no tails on abdomen, long antenna, wings
“spring-tail”, forked jumping organ on underside
Invertebrate Classes
Amphipoda
Decapoda
Isopoda
Gastropoda
Oligochaeta
There are several books for identifying aquatic organisms to a finer taxonomic level. Identification books and keys will
be made available to the students in the laboratory.
Lab 7. Alkalinity, Photosynthesis-Irradiance (PI) curve
Objectives:
1. Learn method for alkalinity titration
2. Explore relationship between photosynthesis and light for a planktonic community
Requirements:
1. Show work on at least one alkalinity titration
2. Compare alkalinity values across habitats
15
3.
Plot P-I curve
Alkalinity: The Buffering Capacity of Water
"....the quantity and kinds of compounds present which collectively shift the pH to the alkaline side of neutrality." Wetzel, 1983
What does this mean to living organisms? Most organisms cannot survive in environments where pH <4.5 or >9.5.
Organisms can also influence alkalinity by affecting the quantity of dissolved CO2 through photosynthesis and
respiration.
Primary compounds occurring in nature include: bicarbonates, carbonates, and hydroxides. Other compounds include
borate, silicate, and phosphates. The major buffers in a lake are primarily dependent upon geology of the basin.
Equilibrium Conditions of Inorganic Carbon in Water:
CO2 (gas) + H20 <—> CO3+ CO2 (dissolved)
H2CO3 + CO2 (dissolved) <—> H+ + HCO3
HCO3- <—> H+ + CO32Alkalinity is determined by titrating with a strong acid solution to equivalence point (the point at which the pH begins to
decline at the same rate at which the acid is added). Units are expressed as microequivalents per liter (µeq L-1) or mg L-1
CaCO3.
Alk, µeq L-1 = (normality of acid)*(volume of acid)*(106)
volume of sample
Alk, mg CaCO3 L-1 = same as above divided by 20
pH
>5.2
5.0
4.8
4.6
color
greenish blue
light blue w/ lavender gray
light pink gray w/ bluish cast
light pink
Alkalinity
(From Standard Methods)
The alkalinity of water is its quantitative capacity to neutralize a strong acid to a designated pH. The measured value
may vary significantly with the end point pH used in the determination. Alkalinity is a measure of a gross property of water
and can be interpreted in terms of specific substances only when the chemical composition of the sample is known.
Alkalinity is significant in many uses and treatments of natural and wastewaters. Because the alkalinity of many
surface waters is primarily a function of carbonate, bicarbonate, and hydroxide content, alkalinity is taken as an indication of
concentration of these constituents. Measured values may include contributions from borates, phosphates, or silicates if
present. Alkalinity in excess of alkaline earth concentrations is significant in determining the suitability of water for
irrigation. Alkalinity measurements are used in the interpretation and control of water and wastewater treatment processes.
Raw domestic wastewater has an alkalinity only slightly greater than that of the water supply. Properly operating anaerobic
digesters typically have supernatant alkalinities in the range of 2,000 to 4,000 mg L-1 as CaCO3. For industrial wastes, the
measurement can indicate change in quality if the source of the sample is known to have generally stable levels of alkalinity.
General Discussion
 Principle: Hydroxyl ions present in a sample as a result of dissociation of hydrolysis of solutes are neutralized by
titration with standard acid. The alkalinity thus depends on the end point pH used.
 For samples of low alkalinity (less the 20 mg L-1 CaCO3) use an extrapolation technique based on the near
proportionality of the concentration of hydrogen ions to the excess of titrant beyond the equivalence point. The amount
of standard acid required to lower the pH exactly 0.30 pH unit is carefully measured. Because the change in pH
16

corresponds to an exact doubling of the hydrogen ion concentration, a simple extrapolation can be made to the
equivalence point.
End Points: When the alkalinity of a water is due entirely to hydroxide, carbonate, or bicarbonate content, the pH at the
equivalence point of the titration is determined by the concentration of CO2 present at that stage. The CO2 concentration
depends in turn on the total carbonate species originally in the sample and any losses that may have occurred during the
titration. The following pH values are suggested as the equivalence points for the corresponding alkalinity concentration
as calcium carbonate.
End point pH
Alkalinity mg L-1
30
150
500
silicates, phosphates
known or suspected
industrial waste or complex system






Total
5.1
4.8
4.5
4.5
Phenolphthalein
8.3
8.3
8.3
8.3
3.7
8.3
Interferences: Soaps oily matter, suspended solids, or precipitates may coat the glass electrode and cause a sluggish
response. Allow additional time between titrant additions to let the electrode come to equilibrium. Do no filter, dilute,
concentrate, or alter the sample in any way.
Selection of method: Determine the alkalinity of the sample from the volume of standard acid required to titrate a
portion of the sample to a designated pH taken from the table. Titrate at room temperature with a properly calibrated pH
meter or electrically operated titrator, or color indicators.
Report an alkalinity less than 20 mg L-1 CaCO3 only if it has been determined by the low alkalinity method.
Construct a titration curve for the standardization of reagents.
Color indicators may be used for routine and control titrations in the absence of interfering color and turbidity and for
preliminary titrations to select the sample size and strength of titrant.
Sample size: For low alkalinity method, titrate a 200 mL sample with 0.02 N H2SO4 from a 10 mL buret.
Procedure
1.) Use 50 ml of unknown water sample
2.) Add 5 drops of BGMRed indicator
3.) Titrate to light pink color with 0.02N H2SO4-; titrate slowly, drop by drop, as you should notice a buffer effect and
then rapid color change.
4.) Optional: check the pH.
5.) Calculculate alkalinity using the formula below.
Calculations
 Potentiometric titration to end point pH:


Alkalinity mg L-1 CaCO3 = A*N*50,000
mL sample
where: A= mL standard acid used and N = normality of standard acid
Report the pH of the end point used as follows. "The alkalinity to pH _____ = _____ mg/L CaCO3" and indicate clearly
if this pH corresponds to an inflection point of the titration curve.
Potentiometric titration of low alkalinity:
Total Alkalinity, mg L-1 CaCO3 = (2 B-C)*N*50,000
mL sample
where B=mL of titrant to first recorded pH, C= total mL of titrant to reach pH 0.3 unit lower, and N = normality of
acid.
17
Precision and Accuracy
No general statement can be made about precision because of the great variation in sample characteristics. The
precision of the titration is likely to be much greater than the uncertainties involved in sampling and handling the sample
before analysis. In the range of 10 to 500 mg L-1 where the alkalinity is due entirely to carbonates or bicarbonates, a standard
deviation of 1 mg L-1 can be achieved.
Photosynthesis-Irradiance Curve (P-I Curve)
Procedure
1. Collect water in BOD bottles (60 mL) and incubate for 2 hours at different light intensities
2. Halt photosynthesis by adding 400 µL MnSO4 and 400 µL alkaline iodide reagent
3. Add 400 µL concentrated H2SO4, shake, put into beaker
4. Titrate with 0.005 N Na2S2O3 to pale straw color using 40 ml of sample
5. Add 2-6 mL of starch and continue until blue color first disappears (1 mL titrant = 1 mg O2 L-1)
6. Plot rate of Net Primary Production (NPP) vs. light and report values for Pmax and compensation point. See text for
details of these parameters
**Steps 1-2 the TA does the day before; Steps 3-6 is a class effort.
18
Laboratory 8: Invertebrate Ecology and Island Biogeography
Streams often are a heterogeneous assemblage of microhabitats that are utilized by many organisms, including
macroinvertebrates. Benthic macroinvertebrates or benthos, have been generally classified as any invertebrate species larger
than 1 mm and occupying the bottom stream substrate (Hynes 1970). Benthos of the Flint Hills region are dominated by
insect, crustacean, annelid, and molluscan fauna. Macroinvertebrates are important in the processing and transport of organic
matter downstream (Vannote et al. 1980). Many are specialized to utilize different sizes and/or forms (i.e. fine or coarse) of
organic matter.
In conjunction with different feeding groups, there is a continuum of flow velocities which benthos inhabit. The
turbulence of flowing waters combined with the mixed assemblage of the bottom substrate provides several niche
opportunities for benthos. Essential to the understanding of flow over surfaces is the Reynolds number. Reynolds number
(Re) is the ratio of inertia (momentum) to viscosity (stickiness) and so quantifies the contribution of these forces to flow. The
higher the Reynolds number, typically the higher the velocity and flow rate across a surface. Reynolds number associated
with microhabitat surfaces may highly affect what macroinvertebrate assemblages are persist.
MacArthur and Wilson (1967) investigated immigration and extinction dynamics in mangrove islands. This has
become a very insightful area of ecology called island biogeography. One of the important ideas they presented was that the
larger islands tend to maintain a higher diversity of fauna than the smaller islands. It is understood that in most situations a
larger, isolated area will be able to supply more resources and niches than a smaller area. This principle can be applied to
stream ecology. If one thinks of rocks (or cobbles) as different islands, there should theoretically be higher diversity on the
larger rocks.
Objectives:
1. Sample and identify aquatic macroinvertebrates from different sized cobble at Pillsbury Crossing.
2. Calculate and compare species richness, Shannon diversity, and evenness for macroinvertebrates collected
3. Comprehend and apply the Island Biogeography Theory to results in a short discussion
Requirements:
1. Calculate Shannon diversity indices (H') and evenness (J') for all samples. Plot the data as a scatter of points on a twodimensional graph and draw a line corresponding to the regression equation. Calculate a simple linear regression using
Species richness, H’, and J’ as dependent (explanatory) variables and rock area as independent (response) variables.
Define your null and alternative hypotheses and conclusions (See explanations below).
2. What other factors, besides larger flow gradients do larger "islands" offer to support a higher macroinvertebrate
diversity? (Hint: What other functions could larger cobble perform that smaller ones do not?). Report logical, biological
explanations for the results you find.
3. Write a short scientific report describing this lab exercise. Include a short introduction describing background
information, methods used so that it is repeatable by others (but not too detailed), the results, and your conclusions. A
well-written report should be concise, so your report should be 1-2 pages, double-spaced, Times New Roman, 12 pt
font. This is not including figures which should be listed after your report.
Procedure:
The class will be divided into groups (2-3 per group). Each group will be responsible for collecting two cobbles
from each of three size classes (small, medium, & large) from riffles at Pillsbury Crossing. Groups should try to select
flattened limestone rocks (this will make surface area measurements more accurate). For the small-medium size classes, the
cobble should be immediately placed into a zip-lock bag and rinsed with a small aliquot of 85% EtOH and stream water.
The large size class will be placed into either a sieve or a white pan, where the invertebrates and surface film (algae, moss,
etc.) will be brushed with a toothbrush. The material should then be placed in a ziplock bag and rinsed with 85% EtOH. The
large cobble will be labeled with corresponding invertebrate samples by tying a string around it and applying a label to the
string.
In the lab, 1/3 of the members will be responsible for measuring the surface area of the rock. To get the surface area,
photograph the rock, label, and small ruler inside a picture. Import the photo into ImageJ (free software, download online).
Set the scale of the photo by setting the picture a known distance from the ruler. Then using the freeform tool, outline the
rock. The other 2/3of the class will be responsible for identifying and counting the invertebrates under a dissecting scope.
Analysis should include data from all members of the class. Use the accompanying data sheets to record data.
Shannon Diversity Index (H')
19
This diversity index takes both the number of taxonomic groups (richness) into account and the spread (or evenness) of the
density over the richness. The larger the H' value, the more diverse the community. It is important to remember that H'
values alone don't mean anything. They are only useful when making comparisons between communities.
How to calculate H':
H' = -∑ (pi * log pi)
where: pi = ni / N
ni = number of individuals within a taxonomic group
N = total number of individuals per sample (rock)
Evenness (J')
As mentioned above evenness describes the spread or aggregation of species in a community.
How to calculate J':
J' = H' / H'max
where: H'max = log s
s = richness or number of taxonomic groups per sample (rock)
Statistical Analysis
Testing for statistical significance is used to determine what the probability is that the affect we are seeing is a real
relationship that exists or from random chance. In other words, we need to test if it was by random chance that we collected
rocks that showed a positive b within a population that actually had b < 0 (explanation of b below).
1.
First, we need to define the null hypothesis (H0) and the alternative hypothesis before we run statistics on collected data.
According to the island biogeography theory, the larger the island, the larger the richness and diversity, so b should be
positive. Based on previous work, we will state the following:
- HA: Species richness, Species Diversity (H’) of macroinvertebrate communities will increase as cobble size
increases.
- Can you state what H0 should be?
2.
The regression equation. There arefour variables in this study: rock size and species richness, species diversity, or
evenness. The rock size is the independent variable (plot on x-axis) and the dependent variable is the species richness,
etc. (plot on y-axis). The regression equation is for a straight line: Y=b + mX. Y is the dependent variable, X is the
independent variable, m and b are the calculated parameters. m is called the regression coefficient or the slope of the
best fit regression line. This tells how much unit change in Y occurs for every unit change in X. The parameter m can
be positive, negative, or zero. For instance, if m > 0, then there is an increase in Y for every increase in X, and if m = 0
then there is no change in Y for a change in X. The parameter b is the Y-intercept or the value of Y when X is zero in
the regression equation.
m=
∑ XY - (∑ X) (∑ Y)
n

∑ X2 - (∑ X)2
n
b can be found by inserting any corresponding X and
Y values from the data
n = the number of samples collected
What to hand in
A report containing:
 Listed sections outline in “Requirements” above.
 Must state whether the HA and H0 are rejected for each hypothesis and regression (how to determine this from statistics
discussed in class) in the Results section.
 Regression statistics, including F-value, DF, R2 value, and P-value in the Results section (theory behind these values will
be discussed in class).
20

All regression statistics (Species Richness, H’, J’) must be reported, so 3 separate regressions (and figures) must be
executed. List your 3 Figures below the body of your report. Remember to label BOTH axes with appropriate units and
include a caption with labels. (Figure 1, Figure 2, etc.). Include graphs generated in Excel or any other statistical
program.
For statistical analysis, you can use Microsoft Excel or R (it’s free! Can download through http://www.r-project.org/ or
http://www.rstudio.com/ for a user friendly version). If interested in R instead of Excel, TA can supply appropriate code. If
you are used to using another statistical package (SAS, etc.), you are welcome to use this also.

Note: You can work together when running statistics, but everyone must complete their own and supply their own output.
21
Lab 9: Introduction to Spectrophotometer and Colorimetric Analyses, SRP, Chlorophyll a
Objectives:
1. Learn general principles behind colorimetric analysis of chemicals
2. Learn principles behind fluorometric analysis of chlorophyll
3. Calculate unknown concentrations from a standard curve
Requirements:
1. Demonstrate an analysis of SRP
2. Demonstrate analysis of chlorophyll and calculation of final concentration
3. Compare SRP and chlorophyll values within and across habitats
Methods used to determine the concentrations of nutrients in aqueous media
 series of standards of known concentration are prepared for analysis
 reagents are added to standards and unknown samples
 color development in the samples is compared with the standards using a spectrophotometer
 plot the absorbance of the standards vs. their concentrations
 concentrations of samples can be determined from graph of standards
Beers Law (1852): fundamental law governing light absorption by a sample (another way to find unknown concentrations
using absorbance values)
A = log (Io/I) = e * c * L
A = absorbance
Io = incident beam (intensity of light going into sample)
I = transmitted beam (intensity of light going out of sample)
e = constant (extinction coefficient, depends on sample)
c = concentration
l = path length
Limitations:
 only holds true for relatively dilute samples (A< 1.5, best when 0.1 < A < 1.0)
 reaction must be at equilibrium (enough time to ensure color change)
 measures absorbance at peak wavelength (must know for specific chemical testing)
Spectrophotometer/Colorimeter: measures amount of light of a particular wavelength that is absorbed by molecules, atoms,
or ions in a sample
Alternative methods for measuring nutrients in water samples include ion selective electrodes (good only for high
concentrations) and ion chromatography.
Soluble Reactive Phosphorous
Precision
Level
Standard Deviation
Operating Range
100µg P L-1
1 µg P L-1
5 - 250 µg P L-1
Sample Condition
 filtered through Whatman GF/C
 stored at 5°C
 Unacidified
 Less than 5 µg AsO4-3 L-1
Method Principle
22



Phosphate, silicate, arsenate and germanate ions react under acidic condition with molybdate to form heteropoly acids
which can be converted, by suitable reducing agents, to blue compounds of uncertain composition
Using appropriate acid and molybdate strength, ascorbic acid as reductant and antimony as a color enhancing species, an
intensely blue colored complex is formed with PO43- and AsO4-3 having an absorbance maximum at 885 nm
It should be noted that, while the formation of the blue complex is specific to PO43-, the reaction conditions are capable
of hydrolysing labile organic phosphorus compounds. This would give an overestimate of PO4-P and hence an
unreliable estimate of biologically available phosphorous.
Reagents
 Acid molybdate antimony: Mix 500mL distilled water, 7.5 g ammonium paramolybdate ((NH4)6Mo7O24. 4 H20), 0.14 g
antimony potassium tartrate, 88 mL conc. sulphuric acid. Cool and dilute to 1000 mL. Keep in dark glass bottle.
 Ascorbic acid: Dissolve 2.5 g of L-ascorbic acid in 100mL of distilled water. This reagent is stable for a few days if kept
refrigerated.
 Mixed molybdate for natural color determination: mix 4 parts of acid molybdate antimony with 1 part distilled water.
 Mixed molybdate for orthophosphate determination: mix 4 parts of acid molybdate antimony with 1 part ascorbic acid.
Stable for 1 day.
 5 mM Phosphate standard: Dissolve 0.6805 g potassium dihydrogen phosphate (KH2PO4) in 100 mL of water and make
to 1000 mL with water..
Procedure
1. Prepare in duplicate 10 mL of orthophosphate standards having a concentration approximating that of samples to be
analyzed.
Standard
µL of 50 µg/mL stock
Conc. Standard (µmol/ L) PO43- -P
1
0
0
2
5
25
3
10
50
4
20
100
5
50
250
6
100
500
2.
3.
Place 10 mL of each standard or 10 mL of distilled water into test tubes. Add 2.0 mL of mixed molybdate reagent for
orthophosphate determination to all tubes.
After 5 minutes and within 3 hours measure absorbance at 885 nm using a 1 cm path length
Calculations Plot concentration versus absorbance for standards, making a standard curve by hand. Use relationship to
determine concentration of samples from absorbance. Alternatively, use linear regression to establish a relationship with
concentration as the dependent variable and absorbance as the independent variable from the standards. Use this equation to
calculate concentration from absorbance for samples.
Chlorophyll a Analysis
Objectives:
Chlorophyll Extraction Method: used to quantify primary production (phytoplankton) from bulk water samples
***ALL CHLOROPHYLL EXTRACTION AND ANALYSIS SHOULD BE DONE IN A DIMLY LIT ROOM AND
SOLUTIONS SHOULD BE KEPT COLD. LIGHT AND HIGH TEMPERATURES DEGRADE CHLOROPHYLL***
Procedure
1. Filter sample through GF/C filters, note volume filtered and place in dark freezer (use filtered samples from field trips)
2. Place filter in centrifuge tube with a known amount of 95% ethanol (EtOH)
3. Place tubes in a 78°C water bath for 5 minutes
4. Cover tubes with aluminum foil to keep out light and set in refrigerator for 24 hours
5. Centrifuge samples (Only if large amount of sediment is present)
6. If samples are visibly green, set fluorometer window to 3X, if not, set window to 10X, if samples are dark green, dilute
to a known volume
7. Zero fluorometer with 95% EtOH in a cuvette
8. Pipette 4 mL of sample into one cuvette (w/o filter fragments)
9. Put cuvette in meter and look at reading:
a. if on 3X window and reading is <40 got to 10X window, rezero meter
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b.
c.
and try again if still <40 by going to the 30X window
If on 30X window and reading <40 don't worry about it
use calibration curves provided in lab to calculate amount of chlorophyll per unit volume
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Lab 10: Lab tour, autoanalyzer, NH4+ assay
Objectives
1. Be able to describe the autoanalyzer.
2. Analyze NH4+ in the samples collected from field sites.
Requirements:
1. Lab tour notes
2. Analysis results
3. Interpret chemistry results
NH4+-N and NO3--N Analysis
NO3--N Analysis -- Cadmium Reduction Method:
 Actually measures both NO2 and NO3
 Sample is passed through a cadmium column which reduces NO3 to NO2
 The reagents react with NO2 to form magenta-like color which is measured with a spectrophotometer
 Interference by Fe, Cu, or other metals is eliminated by a complexing reagent (EDTA)
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This analysis will be demonstrated in the LTER lab on an Autoanalyzer using a program called The Flow Solution by
ALPKEM, the method is the same, but the machine is automated.

Method for Ammonium (NH4+) Determination
Modified to a 10mL sample size from Solorzano (Limnol Oceanogr. 1969. 1:799-801).
Use the smallest test tubes available that can accommodate a 10mL sample. All glassware should be well scrubbed with hot
water, rinsed with 0.1 N HCL, rinsed 6 times with purified water, and rinsed once with reagent grade water. Many different
things can contaminate the assay including smoking, ammonia detergents, freshly cut grass, opening a bottle of Ammonium
hydroxide in the laboratory, etc. Exercise extreme caution. Precise measurements for additions is imperative!!!
Stock Reagents
1. 0.5mM NH+ stock: Take about 1g NH4Cl and dry for 12 h at 100 °C. Make up stock in an acid cleaned 100 mL
volumetric flask. Weigh exactly 0.0535 g of NH4Cl into the flask, add reagent grade water to the 1000 mL mark (read
from bottom of the meniscus). Put in a clean bottle, and label with the solution, date and your initials.
2. Phenol-alcohol: Add 10g of phenol to 100 mL 95% ethyl alcohol.
3. Sodium nitroprusside (nitroferricyanide): Dissolve 1 g in 200 mL of water. Store in dark bottle for not more than 1
month.
4. Alkaline solution: Dissolve 100 g of sodium citrate and 5 g of sodium hydroxide in 500 mL water.
5. Sodium hypochlorite: Use commercial bleach, as new as possible
6. Oxidizing solution: Add 40 mL alkaline solution (4) to 10 mL sodium hypochlorite (5). Make fresh each day.
Assay
Preparation of Standard
Add exactly 10 mL of reagent grade water to each of the 14 test tubes that compose the concentrations for the standard curve.
Add as follows for standard concentrations, duplicate each (to avoid contamination of 0.5uM stock, pour a small amount into
an acid washed beaker):
uL of 0.5 mM stock
0
2
4
8
16
32
64
final conc. NH4+ (uM)
0.0
0.1
0.2
0.4
0.8
1.6
3.2
Samples
Add 10 mL of each filtered sampled to each sample test tube (in duplicate). For each new type of water (i.e. new lake or new
season) make triplicate recovery tubes. To each recovery tube add 25 uL of 0.5 mM stock.
Addition of Reagents
To each tube, add 0.4 mL (400 uL) of phenol solution (2), 0.4 mL nitroferricyanide (3), and 1 mL oxidizing reagent (6). Mix
well (invert screw cap tubes). Let develop for at least 1 h (3 h is better) in darkness. Mix samples while development is
occurring at least once an hour.
Reading samples
Enter appropriate data into spectrophotometer (standard concentrations). For samples with evident blue color, use the
automatic sipper (power machine off before installation). For mildly low levels use 1cm cells, and 5 cm cells for very low
level samples. Set spectrophotometer to 640 nm, put reagent grade water into both cuvettes and auto zero the machine. Read
all of the standards to make a standard curve. Read concentrations. Print standard graph after unknowns are read. Check for
adequate recovery.
Dispose of all assay waste into a labeled bottle. (Label bottle as "ammonium assay waste Phenol 4g/L and Na
nitroferricyanide 0.1 g/L.") Wash tubes with hot water and fill with 0.1 N HCl for storage. This acid can be saved between
assays.
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Lab 11: Tour sewage treatment and water treatment
Objectives
1. Understand basics of how sewage and drinking water are treated.
Requirements:
1. Fieldtrip notes
2. Describe organisms
3. Report and interpret chemistry results (SRP, NO3, NH4, alkalinity)
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References
Allan JD. 1995. Stream Ecology: Structure and Function of Running Waters. Chapman & Hall, NY. 388pp.
Benson, B.B., & D. Krause, Jr. 1984. The concentration and isotopic fractionation of oxygen dissolved in freshwater and
seawater in equilibrium with the atmosphere. Limnology and Oceanography 29: 620.
Brower JE, JH Zar, & CN von Ende. 1990. Field and Laboratory Methods for General Ecology. 3rd ed. Wm. C. Brown
Publishers, Dubuque, IA. 237 pp
Erman DC & NA Erman. 1984. The response of stream macroinvertebrates to substrate size and heterogeneity.
Hydrobiologia. 108:75-82.
Hart DD. 1978. Diversity in stream insects: regulation by rock size and microspatial complexity. Verh. Int. Ver. Theor.
Ang. Limnol. 20:1376-1381.
Hynes HBN. 1970. The Ecology of Running Waters. University of Toronto Press, Toronto. 555 pp.
Larmon TE & IM Henley. 1955. Determination of low alkalinity or acidity in water. Anal. Chem. 27:851.
MacArthur RH & EO Wilson. 1967. The Theory of Island Biogeography. Princeton University Press. 203 pp.
Malmqvist BL, LM Nilsson, & BS Svensson. 1978. Dynamics of detritus in a small stream in southern Sweden and its
influence on the distribution of the bottom animal communities. Oikos 31:3-16.
Merrit RW & KW Cummins (eds.). 1995. An Introduction to the Aquatic Insects of North America. 3rd ed. Kendall/Hunt,
Dubuque, IA.
Murphy J & JP Riley. 1962, A modified single solution method for the determination of phosphate in natural waters. Anal.
Chim. Acta. 27:31-36
Pohland FG & DF Bloodgood. 1963. Laboratory studies on mesophilic and thermophilic anaerobic sludge digestion. J.
Water Pollut. Control. Red. 35:11.
Reice SR. 1980. The role of substratum in benthic macroinvertebrate microdistribution and litter decomposition in a
woodland stream. Ecology 61:580-590.
Shelly TE. 1979. The effect of rock size upon the distribution of species of Orthocladiinae (Chironomidae: Diptera) and
Baetis intercalaris McDunnough (Baetidae: Ephemeroptera). Ecological Entomology 4:95-100.
Ulfstrand S. 1967. Microdistribution of benthic insects (Ephemeroptera, Plecoptera, Trichoptera, Diptera: Simuliidae) in
Lapland streams. Oikos 18:293-310.
Thomas JFJ & JJ Lynch. 1960. Determination of carbonate alkalinity in natural waters. J. Amer. Water Works Assoc.
52:259.
Vogel S. 1988. Life's Devices. Princeton University Press. 367 pp.
Vogel S. 1994. Life in Moving Fluids. 2nd ed. Princeton University Press. 467 pp.
Williams DD & JH Mundie. 1978. Substrate size selection by stream invertebrates and the influence of sand. Limnology &
Oceanography 23:1030-1033.
Zar JH. 1984. Biostatistical Analysis. 2nd ed. Prentice Hall, Englewood Cliffs, NJ. 718 pp.
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