Kansas State University
Advanced Aquatic Ecology (BIOL890)
Methods Manual for Aquatic Ecological
Research:
Long-term monitoring of Pottawattomie State Lake II
and Kings Creek, Kansas.
Jan 2000 version
Authors listed in alphabetical order using last name:
Bernot, R., T. Horton, J. Jeffrey, M. Kemp, M. Quist, S. Schrank,
D. Stagliano, and C. Zachary.
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Table of Contents
Page
Rationale.......................................................................... 3
Study Sites....................................................................... 4
Methods (Pott.St.Lake II)............................................... 4
Methods (Kings Creek)................................................... 10
Recommendations for Future Methods.......................... 12
References....................................................................... 13
Appendix 1 (Species Lists).............................................
Appendix 2 (Zooplankton Images).................................
Appendix 3 (Datasheets)................................................
Appendix 4 (Pott.St.Lake II Equip. List)......................
Appendix 5 (Kings Creek Equip. List ).........................
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17
18
19
23
Figure 1 (Pott.St.Lake II)..............................................
Figure 2 (Kings Creek)..................................................
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26
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Rationale
This manual presents methods developed for addressing factors influencing aquatic ecosystem dynamics. Data
obtained from methods described herein are important for understanding aquatic ecological processes, and for
responsible resource management and conservation. In order to monitor system variability over time, we focus on a
variety of abiotic and biotic parameters that can be measured repeatedly at specified sites. This approach provides
documentation of short-term system fluctuations, at seasonal scale, and also initiates integrated data collection for longterm system analysis, decade to century scale.
Due to the complexity of physicochemical processes and biotic interactions, we feel it is important to track
system characteristics at several levels rather than just a few. A multifaceted research approach will enable investigators
to monitor trends over space and time, establish biotic inventories, quantify environmental impacts and ecosystem health,
ascertain stream reference sites, and provide baseline data supporting future research and analyses. Our ultimate goal is
to conduct research that has the capacity to gain a greater understanding of aquatic ecosystem dynamics, and
simultaneously educate future graduate students. In simple terms, our research design measures one lentic and one lotic
system during the spring and fall seasons. Four levels of system monitoring were selected, justification for each follows.
Physical & Chemical Parameters. – 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, O 2 availability, temperature, and pH can control
biotic communities. These factors can also indicate anthropogenic effects on the system. Physical properties of water
make it especially vulnerable to pollution inputs. Such inputs can alter the nutrients available and therefore change
energy flow within aquatic systems. Water pollution can cause eutrophication, methelhemoglobemia, and toxic algal
blooms. Thus, it is essential to maintain an active record of the physical and chemical parameters of water bodies.
Plankton. – The plankton community of temperate freshwater lakes varies significantly from year to year as well as
from season to season (Lampert and Sommer 1997). Also, vertical structuring of the community at any one time is
likely during periods of lake stratification (Leibold and Tessier 1991). Many factors such as resource quantity and
quality, food web structure, and physical parameters may ultimately influence these dynamics. Therefore, long-term
monitoring of plankton in concert with other biota and abiotic measures, may elucidate dominant mechanisms associated
with lentic dynamics. The instantaneous vertical distribution of plankton, before and after summer periods, may also be
important for understanding seasonal linkages to other variables.
Macroinvertebrates. – Benthic macroinvertebrates dominate the biomass in the sediments of most freshwaters and
represent an important component of productivity (Downing 1984). In lentic systems, some of the important
relationships among the benthic organisms involve comparisons of the littoral zone with the profundal (Wetzel 1983).
The littoral zone typically has a more productive and heterogeneous aquatic community (Wetzel 1983), mainly because
of aquatic macrophytes. Aquatic macrophytes support higher invertebrate diversity and abundance when compared to
adjacent non-vegetated zones (Dvorak & Best, 1982; Iversen et al., 1985).
In lotic systems, macroinvertebrates show spatiotemporal variability, are relatively immobile, ubiquitous,
highly diverse in species, and have relatively long life cycles. Sampling can be conducted with simple and inexpensive
equipment, the taxonomy of many groups is well documented, data analysis methods have been developed and widely
used, and pollution responses of many species have been established (Rosenberg and Resh 1993). Therefore,
macroinvertebrates have been shown to be effective bioindicators of system impairment (Resh 1995). The
macroinvertebrate component of aquatic systems is often considered a contributing force in nutrient cycling, and also a
substantial link between primary producers and higher trophic levels.
Fish. -- As a whole, fishes are important consumers at every trophic level and can dominate some aquatic food webs.
Fishes can be an essential food base for terrestrial mammals, avifauna, and herpetofauna. Physicochemical dimensions
and both plankton and macroinvertebrate communities may vary in a way that influence fish communities by changing
environmental suitability and food resources. The long-term monitoring approach taken here will aid us in
understanding the structure and dynamics of fish communities, and their role in aquatic ecosystem function.
Study Sites
The lentic site selected for this project was Pottawattomie State Lake II. This lake is located in Pottawattomie
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county, Kansas, owned by the Kansas Department of Wildlife and Parks (KDWP), and managed for camping and sport
fishing (Figure 1). The lake’s watershed drains tallgrass prairie rangeland and mixed hardwood forest of its tributaries.
The Konza Prairie Research Natural Area (KPRNA) is a 3,487 ha site located in central Geary and Riley
counties, Kansas. KPRNA is owned by the Nature Conservancy, managed by Kansas State University-Division of
Biology, and part of the National Science Foundation Long-Term Ecological Research Program. Greater than 90% of
the site is tallgrass prairie with limited woodlands along streams. These streams drain large prairie watersheds and
exhibit unstable flow regimes and harsh fluctuations in environmental conditions. The lotic site selected for this project
was King’s Creek, located on KPRNA, and the permanent study reach lies mid-catchment with a riparian mixed
hardwood forest. This stream reach is 100-m in length and is upstream of the public nature trail crossing (Figure 2).
Methods (Part 1 of 2)- Pottawatomie State Lake II
Aside from fish sample sites, all other samples will be taken from three fixed stations (stations 1, 2, and 3; Figure 1).
Station 1 represents the deepest point in the lake, and stations 2 and 3 are located in the two inundated tributaries, “creek
arms”.
Physical & Chemical Characteristics. – Several parameters will be measured at the established three sites, at one meter
intervals within the water column, to determine physicochemical characteristics of the lake. A Hydrolab will be used to
measure dissolved oxygen, temperature, redox potential, conductivity, and pH. Other variables measured include light
attenuation and NH4, NO3, PO4, SRP, TN, and TP (methods described below).
Water samples for determining NH4-N, NO3-N, PO4-P, SRP, TN, and TP will be taken using a Van Dorn Bottle. The
Van Dorn Sampler has a cord that is marked in ½ meter increments. Two samples will be taken every meter.
1.
2.
3.
4.
Hook suction cups on top.
Lower device to appropriate depth.
Drop messenger and retrieve Van Dorn Bottle.
Pour water sample into Nalgene bottle, allow bottle to overflow with the sample water three times.
5.
Filter 100 ml of a sample and label as filtered or unfiltered, depth, date, initials, and site (use GF/F
filters).
Samples are to be kept cool until analysis can be done.
6.
All chemical analyses, except for the NH4 analysis, are performed on a Technicon auto-analyzer. The NH4 analysis will
be done by hand using the phenol-hypochlorite method and a Hitachi double-beam spectrophotometer (Solarzano, 1974,
see Standard Methods of Water and Wastewater Analysis).
Light measurements will be made in two ways. First, we will use a Secchi disk to make an indirect measurement of
water clarity.
1.
2.
Lower Secchi into the water on the shady calm side of the boat until it can just, barely be detected.
Record the depth of the disk.
The second method is a more direct measurement of light allowing determination of depth at which photoinhibition
occurs. A LiCor photometer will be used to measure the available light in mol quanta/m 2/sec every meter with depth. It
is necessary to take the light readings on a clear day with no clouds.
1.
2.
Take a light measurement above water in open air.
To take a measurement, press “on” and lowering the photometer to the required depth, the “hold”
button is pressed down while the measurement is read.
Plankton. – The composite biomass of phytoplankton populations will be estimated by measuring the concentration of
chlorophyll a pigments in water samples collected from depths at one-meter intervals.
Phytoplankton collection – at 1m depth intervals
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1.
2.
3.
4.
Lower Van Dorn to desired depth and send messenger to trap water sample.
Filter 100mL of sample water through Whatman GF/F filter (discard unfiltered).
Fold filter in half (green on the inside) and wrap in aluminum foil and label.
Begin extraction procedure in lab or immediately freeze, and analyze within 2 wks.
Extraction – Extraction of chlorophyll a pigments will follow modified procedures of Standard Methods for Examination
of Water and Wastewater (1998) and Sartory and Grobbelaar (1984).
Note: extracting and measuring chlorophyll a should be done in subdued lighting (lights off!) to prevent pigment
degradation.
1.
2.
3.
Place filter in a test tube with 3.5mL of 90% ethanol.
Boil for 5 minutes at 79-80C.
Cover all the samples with aluminum foil (to keep them in the dark), and place in the refrigerator for
12-20 hours.
Fluorometric Method
1.
2.
3.
Centrifuge samples until all the pieces of the filter are on the bottom of the test tube. Use micropipette
to carefully pipette liquid into fluorometer tube.
Measure sample fluorescence at a sensitivity setting (3x, 10x, or 30x) that will provide a midscale
reading.
Convert fluorescence readings to concentrations of chlorophyll a (mg/L) by using the appropriate
regression equations below:
where:
y = fluorometer reading
x = chlorophyll a concentration (mg/L)
3x:
10x:
30x:
1x:
y = 220.1721x – 0.79579
y = 788.6752x – 0.8809
y = 1884.286x – 0.11786
y = 81.07368x + 0.742807
Zooplankton collection – at 1m depth intervals
1.
2.
3.
4.
Slowly lower Schindler sampler to desired depth, then raise sampler to the surface. (12 L of water
should be trapped from the desired depth only).
Lift sampler out of the water so that water drains from the mesh netting only.
Pour sample from catch container into sampling cup (rinse with 70% ethanol).
Close sampling cup and label with depth, date, initials, and site.
Zooplankton Enumeration – modified from Wetzel and Likens (1991)
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
Split sample if it is “too dense” (i.e. >500 organisms). To split sample:
Use ring stand to hold splitter.
Place a beaker under each splitter tube.
Pour entire sample into top.
Wash remaining debris with wash bottle.
Repeat as needed.
Bring sample to 100 mL volume in graduated cylinder.
If already >100 mL, settle for 15 minutes, and pipette off excess H20.
Bring sample to 90mL, pour into beaker, then use additional 10mL to wash remaining debris into
beaker (total 100 mL).
Stir sample in beaker with volumetric pipette (with 5 mL attachment) for 15 seconds.
Take subsample (5 mL) with pipette.
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12.
13.
14.
15.
16.
Pour subsample into counting wheel groove using glass funnel held by ringstand.
Wash remaining debris from pipette or funnel into groove.
Let sample settle into groove for a minute or two (note: large amounts of algae may require more
settling time).
Use dissecting microscope at 12x to count and identify all organisms in groove.
Rotate disk slowly and record each organism using a counter.
Report zooplankton as number per liter:
# of zoops / L = C x V’
V” x V”’
where:
C = # of organisms counted
V’ = volume of the concentrated sample (L)
V” = volume counted (L)
V”’ = volume of the grab sample (L) = 12 L
Macroinvertebrates. – Station 1 will not be sampled for macrophytic invertebrates because of its location near the dam
which generally lacks macrophytes.
From each fixed station, a transect will be established perpendicular to the shoreline. The distance to the shore from the
fixed station divided by 3 will mark two sampling sites in addition to the fixed site (marker buoys used for sites 2 and 3).
One benthic sample will be collected for each site. At sites 2 and 3, semi-quantitative macrophyte sweeps will be taken
along the transect when macrophytes become apparent. Invertebrate samples will be taken after physicochemical
measurements and plankton collections are completed.
Note: In lab, before sampling, mix up a gallon of 10 % buffered formalin. Pour 185 ml of 37 % Formaldehyde into a
1000 ml graduated-cylinder and fill with water. Repeat four times. Add 5 grams CaCO 3 / liter of 10 % formalin=20
grams. Add ~1/4 teaspoon of Phloxine B and shake.
Benthic Sample: Quantitative estimates of macroinvertebrate density and biomass will be assessed with an Ekman grab
sampler.
1.
2.
3.
4.
5.
Lower Ekman to the bottom, keep line taut (note depth on the cord).
Send messenger down to spring jaws.
As the Ekman is retrieved, a bucket will be placed under it at the water surface to avoid losing contents
(if jaws did not shut fully, retake sample)
Release sample into the bucket, and rinse the inside of the Ekman.
Pour onto a 250µm standard sieve; this may take several washes if clogging occurs. A steady flow of
water will usually unclog mesh, but tapping the bottom of the sieve often helps.
Work the sample to one side of the sieve and wash into an insect pollination bag (poly-bag). Add label and preserve
with 10 % buffered formalin stained with Phloxine B. Squeeze bag to remove air, twist and tie an overhand knot. Place
in another poly-bag. Labels should include site number, date, Eckman number, depth and initials.
Macrophyte Sample: Timed macrophyte sweep samples will be taken with a dipnet to assess the relative abundance and
diversity of epiphytic macroinvertebrates in the littoral zone. Two sweeps will be made for each site.
1.
2.
3.
A standard (46.5 x 24.5 cm) rectangular dipnet (500µm mesh size) will be forced through the
vegetation in a sweeping motion for a period of 30 seconds.
Place all macrophytes hanging off the sides into the net and swish water around the net bag to
consolidate sample into a corner of the net.
Place net contents (dislodged organisms and epiphytes) into a poly-bag, and examine net mesh for
remaining organisms. Label and preserve with formalin.
Benthic Organism Separation, Enumeration, and Identification:
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1.
2.
3.
4.
Open poly-bag, fill with water and remove label.
Pour sample into bucket and further dilute with water
Pour contents onto nested (1mm and 250µm) sieves, wash sample contents until clean (no more silt
going down drain).
Separate sieves and rinse contents of 1mm into a white-bottom enamel pan.
Bring pan to an illuminator, remove all organisms with forceps and place them into a scintillation vial filled with 70 %
EtOH. Double-check pan for organisms under the dissecting scope. Label vial with site number, date, Eckman replicate,
size fraction, subsample fraction, and depth.
Examine detritus on 250µm sieve, and if loaded with insects (>200) or too much volume (judgement call, but usually if
more than a tablespoon) then split sample with the Zooplankton splitter (see Zooplankton methods).
1.
2.
3.
4.
Place subsample in a petri dish and pick all organisms (incl. zooplankton) under the microscope (1020x) into scintillation vial w/ 70% EtOH.
Identify organisms to the lowest taxon possible & count.
For the subsample, multiply the # of each individual taxon by the subsample denominator (ex. 50
chironomids in a ¼ sample =200 for the <1mm fraction)
Combine the <1mm subsample total w/ the >1mm total per taxa & multiply
by 43.3 (Ekman area/m2) to get #/m2. To continue with the chironomid example:
# of chironomids/m2 =(50 in >1mm + 200 in <1mm) x 43.3= 10,825
Macrophytic Invertebrate Separation, Enumeration, and Identification:
1.
2.
3.
4.
5.
6.
Open poly-bag, fill with water and remove label.
Pour sample into bucket and further dilute with water, agitate macrophytes.
Pour contents onto 500µm sieve (retaining large macrophytes until final rinse), wash sample contents
until clean (no more silt going down drain), may take several rinses.
Spread macrophytes out in a large tray and examine for invertebrates.
Rinse contents of 500µm sieve into a white-bottom enamel pan and pick in proportion of abundance
(pick all obvious or large/rare taxa and then the numerous taxa) with a goal of about 200 organisms.
Place in a scintillation vial with 70% EtOH and label with site number, method and number, date and
position sample was taken from.
Fish. – Fish samples will be collected using methods and fixed sampling locations defined by KDWP protocols (ref
here). A boat-mounted Coffelt electrofishing system with pulsed direct current (DC) will be used for fish collection.
Field size and strength will be standardized for each sampling period. A total of one hour of actual electrofishing time
will be conducted during each sampling period (i.e., spring and fall), beginning 0.5 hours after sunset. After each 15
minutes of electrofishing, fish will be brought to shore and processed (see below).
Gill nets and trap nets will be set from sunset to sunrise and should not be deployed longer than 24 hours. Nets
will be set perpendicular to shore and trap nets should be positioned with the frame in an upright position. The gill net
complement (i.e., 1", 1.5", and 2.5") and each trap net represent one unit of effort each.
All fish will be processed as soon as possible after collection. Each fish will be identified to species and
measured to the nearest mm (total length). Weights will be recorded to the nearest gram from 10 fish per centimeter
length group per species. Ten or more scales will be removed from target species at the tip of the pectoral fin and below
the lateral line (Figure 16.2; DeVries and Frie 1996). Scale samples will be taken from 10 stock-length fish per
centimeter length group for each target species; largemouth bass Micropterus salmoides, bluegill Lepomis macrochirus,
redear sunfish Lepomis microlophus, warmouth Lepomis gulosis, white crappie Pomoxis annularis, and black crappie
Pomoxis nigromaculatus.
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Methods (Part 2 of 2)- Kings Creek
Water samples are taken three times a week by the LTER program at the public nature trail crossing. Any information
regarding the chemical concentrations of the stream can be obtained by contacting Walter Dodds (Bushnell Hall).
Macrohabitat. – Each macrohabitat (i.e., pool, riffle, run) will have three transects set perpendicular to stream-flow.
Transects will be placed at 1/4, ½ , and 3/4 of macrohabitat length. Three widths (nearest decimeter; one per transect)
and one length (m) for each macrohabitat will be measured with a rangefinder (Impulse 2). Any logs, boulders, or debris
should be included in the width measurement; however, accumulations of inorganic sediments greater than 0.31 m in
width will be considered islands and should not be included in the stream width measurement (Platts et al. 1983).
Current velocity (m/s), depth (m), and substrate type, will be measured at 20, 40, 50, 60, and 80% of the stream width at
each transect (Platts et al. 1983). Current velocity will be measured using a Marsh-McBirney Flo-Mate Model 2000,
flowmeter. At depths less than 0.75 m, one measurement of the velocity will be taken at 60% of the water depth
(Buchanan and Somers 1969; McMahon et al. 1996). At depths greater than 0.75 m, velocity will be measured at 20%
and 80% of the water depth (Buchanan and Somers 1969). Stream depth will be measured using a calibrated top-set
wading rod to the nearest centimeter. Substrate will be measured using the point transect method and will be classified
using a modified Wentworth system (McMahon et al. 1996), except for the inclusion of a bedrock category and the
pooling of sand categories. In areas where visual examination of substrate is hindered by turbidity or depth, substrate
particle size will be estimated by touch with the wading rod (Platts et al. 1983). Substrate categories are as follows:
bedrock or boulder (> 256 mm), cobble (65 - 256 mm), pebble (32 - 64 mm), gravel (2 - 32 mm), sand (0.0625 - 2 mm),
silt (0.0039 - 0.0625 mm), clay (< 0.0039 mm).
Mesohabitat. – The area of mesohabitats (m2) will be measured for each macrohabitat. Mesohabitat categories include:
log, log complex, aquatic vegetation, brush pile, boulder, overhanging vegetation, and root wad. To be classified as one
of the previously mentioned categories, the habitat in question must have a minimum width and length of 0.1 m. Two
widths and two lengths from each mesohabitat will be recorded. To maintain consistent classification of mesohabitats
the graduate student currently responsible for the long-term study (e.g., Sally Schrank until June 2000) will classify all
mesohabitats in the study reach.
Canopy Cover. – Canopy cover of the stream reach will be determined using a densiometer. At the center of each
macrohabitat, four measurements will be taken: facing downstream, facing upstream, and facing each bank. The
densiometer is held chest-level and the numbers of squares with light (no vegetation) are counted. The percent canopy
cover is then calculated from the following equation:
% cover = average number of squares with light x 96 x 1.04
These directions are also on the back of the densiometer and can be referred to in the field.
Periphyton. – Periphyton biomass will be determined by obtaining the ash-free dry mass (AFDM) and chlorophyll
concentrations of collected samples. This involves drying the collected samples to a constant weight, oxidizing them in a
muffle furnace and reweighing the oxidized samples. The loss in weight upon oxidation is referred to as the AFDM.
Prior to field sampling, filters should be ashed in the muffle furnace (550 C, 3 hours) and their weight recorded. These
pre-weighed filters will be used to filter the biomass samples. Periphyton will be collected by selecting four relatively
flat rocks from the two riffles in the reach and placing the epilithon sampler on the rock to form a sealed section of the
rock. The sealed section is then brushed/agitated to displace all periphyton and then sucked up using a turkey baster and
placed into the sampling container. The rock should be washed and the liquid within the sealed section retrieved several
time to ensure all of the periphyton is retrieved. Furthermore, it is important to rinse the brush into the sampling
container to obtain all of the periphyton left on the bristles.
Two of the rocks from each riffle site will be used for biomass calculations. These samples will be filtered onto
the pre-weighed Whitman GF/C filter (several if necessary) and placed in the drying oven. The samples are weighed,
after thoroughly dried, and placed into the muffle furnace (550 C, 3 hours). Once the samples are ashed, they are re-wet
by gently squirting with water and again allowed to dry in the drying over. Finally, when dried they are again weighed.
Ash free dry mass (AFDM) is obtained using the following calculation:
AFDM = (dry wt – filter wt) – (ash wt – filter wt)
The average AFDM is then scaled up to the whole stream using the area of the epilithon sampler (__area size?).
9
The samples from each of the microhabitats will be taken back to the laboratory and placed into the drying
oven, weighed, and then placed into the muffle furnace and again weighed to determine AFDM. The remaining two
samples from each of the riffles will be used to determine the chlorophyll within the reach. The samples will again be
filtered onto a GF/C (no pre-weighing necessary) and the filters will be folded and wrapped in tin foil. These samples
will be placed in the freezer upon return from the field for future chlorophyll extractions.
Macroinvertebrates. – Macroinvertebrates will be quantitatively sampled in riffles by taking 3 replicate Surber samples.
Qualitative collections will be made with a standard-sized dipnet in all habitats (one composite of 20 jabs in proportion
to riffle, pool, and run availability). Surber samplers should be placed in water at least 12 cm deep and all substrata
should be dislodged and rubbed clean; keep large substrate out of the surber’s bag. Dipnet jabs in pools should focus on
submerged woody debris, roots, undercut banks, and macrophytes. Dipnet jabs in riffles should focus on cobble and
pebble substrate. All dipnet jabs should be approximately 0.5 m in length. All samples will be placed in labeled poly
bags, immediately preserved with Phloxine-B stained 10% formalin, and later sorted and identified to the lowest taxon
possible. These methods follow the updated version of Plafkin et al. (1989). Plastic paper labels should include date,
riffle/pool complex number (RPC 1 or 2), and method (dipnet, surber 1, 2, or 3). In the laboratory, dipnet samples will
be sorted using a two-phase processing method and surber samples will be sorted using a nested-sieve method. For both
methods, a 200 organism fixed-count technique is typically employed, where processing continues until the target
number is achieved. Although a 100 organism count is generally sensitive to species richness and abundance, the power
of discriminating differences among assemblages can be reduced (Barbour and Gerritsen 1996, Courtemanch 1996,
Vinson and Hawkins 1996). Therefore, attempts will be made to obtain 200 organisms for all samples.
A dipnet sample is first washed onto a 500 um sieve and then rinsed into a sorting tray. The two-phase
processing method first searches the entire sample for large and rare organisms, then a subsample based upon a fraction
of the whole is searched. If organisms are few in the first 1/4 of the second phase, a second 1/4 should be processed, and
so on. A surber sample should be washed onto nested 1mm and 250µm sieves. The entire collection from the 1 mm
sieve is first picked, then subsampling of the 250µm sieve follows until obtaining at least 200 organisms (see Pott2
macroinvertebrate section for technique details).
Fish. – The backpack unit should be set on DC current, at 30 cycles per second (cps), between 100-300 volts and 3-5
amps (depending on the conductivity of the water, volts may be adjusted accordingly to obtain 3-5 amps). The generator
output will be set on 300 volts (denoted as 300 VA on backpack unit). Each macrohabitat will be sampled
independently. Electrofishing will begin at the downstream end of each macrohabitat and proceed upstream, ending
when the uppermost portion of the macrohabitat is reached. The anode will be moved in a smooth arc-like motion from
bank to bank. Two assistants will net fish–on either side of the backpack unit–and place them in a bucket that is
approximately half-full of water. Total time electrofishing will be recorded for each macrohabitat (read and reset timer
on the backpack unit–with the switch on the anode–prior to electrofishing each macrohabitat). Fish from each
macrohabitat will be worked up separately.
All fish will be sorted by species and worked up as soon as possible. Total length (mm) will be measured from
the first 100 fish of each species, subsequent fish of that species will be counted. All fish will be kept separate based on
macrohabitat type and number. Species name abbreviations and data sheets should follow standards in Appendix. All
fish will be returned to the macrohabitat from which they were collected.
Recommendations for Future Methods
Due to logistically constraints there are several techniques that we were unable to incorporate into this project,
yet we feel that some may be warranted. For the lentic site, bathymetric mapping would better portray water volume,
depth, and other macrohabitat features. Also for the lentic site, consideration should be given to future faunal surveys for
tributary fish, lake amphibians and reptiles, adult Odonates, and water birds. For the lotic site, the stream channel should
be characterized in terms of other physical features, e.g., in a way that would show channel movement over time (see
Harrelson et al. 1994). Also for the lotic site, aspects of crayfish and amphibian influences on stream system function
should be addressed. Some of these suggestions might be possible if viewed as multi-annual endeavors rather than
mandatory semi-annual surveys; flexible to new students’ specialties.
Implicit to this project endeavor are large data files covering both biotic and abiotic variables for two sites.
Providing a way to have these data widely and easily accessible to users is an objective that will require future
refinement. Having protocols for data archival and backup should be established. Data files for 1999 exist as
10
independent constructions, with designs that are varied. Although these files can be brought into a relational database (a
database of databases), the overall design will lack some interconnectivity necessary for comprehensive queries.
References
Barbour, M.T., and J. Gerritsen. 1996. Subsampling of benthic samples: a defense of the fixed-count method. Journal
of the North American Benthological Society 15(3):386-391.
Buchanan, T. J., and W. D. Somers. 1969. Discharge measurements at gaging stations. United States Geological
Survey, Techniques of Water-Resources Investigations, Book 3, Washington D.C.
Courtemanch, D.L. 1996. Commentary on the subsampling procedures used for rapid bioassessments. Journal of the
North American Benthological Society 15(3):381-385.
Cross, F.B., and J. T. Collins. 1995. Fishes in Kansas, 2 nd edition. University of Kansas, Lawrence.
DeVries, D. R., and R. V. Frie. 1996. Determination of age and growth. Pages __-__ in Murphy, B. R., and D. W.
Willis, editors. 1996. Fisheries techniques, 2 nd edition. American Fisheries Society, Bethesda, Maryland.
Downing, J.A. 1984. Sampling the benthos of standing waters. Pages 87-130 in J.A. Downing and F.H. Rigler, Editors.
A Manual on Methods for the Assessment of Secondary
Productivity. 2 nd edition. Blackwell, Oxford.
Dvorak, J., and E.P. Best, 1982. Macro-invertebrate communities associated with the macrophytes of Lake Vechten:
structural and functional relationships. Hydrobiologia 95: 115-126.
Greenberg, A.E., L.S. Clesceri, and A.D. Eaton. 1998. Standard Methods for the Examination of Water and
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