The objective of this lab exercise is to introduce students to watershed ecology, ecological factors that affect water quality, measurement of water quality parameters, and use of ecological indicators to assess water quality.
Please see the Elements of Ecology textbook for background information about stream and lake ecosystems (pages 518-530 in the Seventh Edition) and the EPA manuals on water quality and macroinvertebrate sampling (see links on Syllabus page of web site).
Freshwater resources are critical for human uses and for the organisms that live in or rely on them. Many human activities have strong effects on water quality, including land use
(e.g., water percolating through a forest soil is effectively filtered, while water running off a paved road dissolves pollutants on that road and transfers them to nearby aquatic ecosystems), use of septic systems (many of which leak), fertilizer and pesticide application, atmospheric pollution and deposition (including ‘acid rain’), and climate change. The term ‘watershed’ is used to refer to the land area from which water drains into a particular stream, river, or lake. Abiotic conditions in aquatic ecosystems impact the organisms these systems harbor directly and therefore control the composition of aquatic communities. While it can be prohibitively expensive to monitor the entire suite of potential pollutants and other abiotic conditions, a few key parameters have a strong influence on biological communities and are relatively easy to monitor. For example, most aquatic ecosystems are limited by nitrogen (typically present as nitrate or ammonium) and/or phosphorous (phosphate). Increasing one or both of these key nutrients can stimulate algal blooms, affecting turbidity, light penetration, oxygen concentrations, and population dynamics of organisms that graze algae (zooplankton), decompose them (bacteria), as well as higher trophic levels. pH and temperature are key abiotic factors that influence community composition and water quality. Similarly, dissolved oxygen affects any organism that respires aerobically, ranging from fish to aerobic bacteria. Dissolved oxygen tends to be very high in streams, as turbulence and movement replenishes oxygen that is consumed by aerobic respiration. However, in deep pools or highly polluted streams, dissolved oxygen concentrations may be very low
(hypoxic) or zero (anoxic).
In the second week of this lab module, we will be analyzing macroinvertebrate communities collected from Great Brook in Week 1. Macroinvertebrate communities serve as ecological indicators of abiotic conditions in aquatic environments. As sessile
(immobile) organisms, they are impacted by highly localized conditions. Because specific types of macroinvertebrates are able to tolerate and/or thrive under different environmental conditions, their presence or absence can serve as indicators of those conditions in a particular location. In addition, while sampling and analysis of water chemistry is typically done by taking discrete samples at a limited number of locations and times, macroinvertebrates remain in their environments for extended periods and

therefore effectively integrate environmental conditions over time. Macroinvertebrates are relatively easy to identify (to the family level); they have a broad range of trophic levels and pollution tolerances, making them a robust means to assess environmental conditions; sampling macroinvertebrates is straightforward and not detrimental to the environment; and databases on their distributions and tolerances for various environmental conditions are already available. Together, these attributes make macroinvertebrate community analysis an effective tool for assessing water quality.
In order to reduce resistance to water current and avoid being swept downstream, many macroinvertebrates have ‘streamlined’ body forms. For example, the larval forms of many insects cling to undersurfaces of stones (e.g., mayflies and stoneflies) or construct protective cases of sand or pebbles and cement them to bottoms of stones (caddisfly larvae). Other macroinvertebrates, such as snails, have a sticky undersurface to cling to and move on stones and rubble. You will be observing these morphologies in week 2 of this lab module.
Macroinvertebrates serve as the primary detritivores in rivers and streams, consuming dead organic matter and aiding in its decomposition. They are typically classified by how they obtain food, as shown in Table 1.
Table 1. Categories of macroinvertebrates based on food acquisition.
Shredders* Feed on particulate organic matter (OM) > 2 mm
Collector-gatherers Dead OM particles from stream
Collector-filterers bed
Sift small particles from water column
Grazer-scrapers
Carnivores
Scrape off and consume OM attached to rocks/stones (e.g., bacteria, algae, fungi, dead material)
Caddis-fly larvae, shrimp,
May-fly larvae, oligochaete worms
Blackfly larvae, caddis fly larvae
Cased caddis, mayfly larvae
Dragon fly larvae, leech, alderfly larvae
*Shredders are actually feeding primarily on bacteria and fungi that colonize dead OM
(i.e., leaves) soon after it enters the stream ecosystem.
Fast-moving streams typically have lower temperatures and higher dissolved oxygen concentrations than slow-moving streams. Macroinvertebrates that inhabit fast- vs. slowmoving streams are shown in Table 2. Similarly, certain macroinvertebrates are more tolerant of polluted conditions, while others are intolerant or facultative (Table 3).
Several indices have been developed that take tolerance/intolerance of pollution into consideration in order to calculate a number that represents the level of pollution in a

particular ecosystem. For example, the percent composition of pollutant-tolerant vs. – intolerant individuals has been found to be correlated with quantitative measures of pollutants. The Hilsenhoff Biotic Index is an index that is oriented towards organic pollutants and takes into consideration organisms sensitive or tolerant to dissolved oxygen levels and/or the concentration of organic compounds. Alternatively, the Biotic
Condition Index is oriented towards the level of sedimentation in aquatic ecosystems. In addition, in general, streams in which pollution is not a major stressor have a higher diversity and abundance of macroinvertebrates (although other environmental factors clearly influence diversity and abundance) than polluted streams.
Table 2. Macroinvertebrates that typically inhabit fast- vs. slow-moving streams.
Fast stream Slow stream blackfly larva dragonfly nymph net-spinning caddisfly water strider stone case caddisfly mayfly nymph stonefly nymph damselfly larvae water boatman bloodworm water penny donbsonfly larvae cranefly larvae crayfish
Table 3. Tolerance of polluted conditions by various macroinvertebrates.
Intolerant Tolerant Facultative
Operculate snails Sludge worms net-spinning caddisfly midge larvae stone case caddisfly leeches mayfly nymph pulmonate snails scuds blackfly larvae stonefly nymph some polycheates riffle beetles
Hellgrammites(dobson fly larvae) most snails sowbugs
In order to describe the diversity of macroinvertebrate communities you sample, you will be determining the species richness and the
Simpson’s diversity index, as well as plotting rank-abundance curves for each sample you collect (2 samples from one location), as well as for two samples from a location sampled by your classmates.
Species richness is simply the total number of taxa observed in a given sample (e.g., if four different species were found, the species richness, or S , = 4). Diversity indices

attempt to combine both species richness and the evenness of their abundances. For example, in two communities with the same number of species, the community that has a more even distribution of species abundances should be considered more diverse. Thus, if community A and B are each comprised of 4 species, their species richness values are both four. However, if the relative abundances of species are 25%, 25%, 25%, and 25% for community A (an even distribution of species abundances), while they are 95%, 3%,
1%, and 1% for community B (uneven), then a diversity index should reflect that the diversity of community A is higher than that of community B. Simpson’s diversity index is a number corresponding to the number of randomly selected pairs of individuals that must be drawn from a community to have an even chance of obtaining a pair with both individuals of the same species. So, if a community has one species, you only have to draw 1 pair (D s
=1), while if every individual in community belongs to a different species, D s
is equal to infinity. The equation for Simpson’s diversity index (D s
) is below:
D s


N ( N

1) n i
( n i

1)
N= Total number of individuals of all species n i
= Number of individuals of a species i
Expressing species richness and evenness in one number provides a simple means for comparing the diversities of many communities. However, because information about abundance and species richness is combined, their individual contributions to diversity are hidden. Rank-abundance curves can be used to reveal this information.
In order to plot a rank-abundance curve, the relative abundances of each taxon must first be determined. An example of data obtained from a macroinvertebrate community analysis is shown below in Table 4. Once the total number of individuals are determined for each taxon ( n i
), the total number of individuals across all taxa for a given sample is simply:
N =

n i .
The relative abundance is determined by dividing n i
by the total number of individuals for the sample, N:
P i
= n i
/N.
Then, each taxon is ranked according to its relative abundance – e.g., the taxon with the highest relative abundance has a rank of 1, that with the second highest relative abundance has a rank of 2, and so on. Relative abundances are then plotted on the y-axis against rank on the x-axis for each community. Thus, communities that are diverse, i.e., have a high species richness and high degree of evenness have rank-abundance curves with slopes near zero (horizontal lines), while those that are not diverse (few species,

uneven distribution) have steep negative slopes. An example of a rank-abundance curve for the data in Table 4 is shown in Figure 1 below.
Table 4. Example data for macroinvertebrate community calculations.
Taxon # individuals (n i
) Relative abundance
(p i
) rank
Mayfly nymph
Caddisfly
Stonefly nymph
50
30
20
Total number of individuals (N) 100
0.5
0.3
0.2
1
2
3
Rank-abundance curve
0.6
0.5
0.4
0.3
0.2
0.1
0
0 1 2
Rank
Figure 1. Example of a rank-abundance curve.
3 4


Hip-waders
 meter sticks (to measure stream depth)
 water samplers
 dissolved oxygen (DO) water samplers
 water analysis kits (DO, NO
3
, PO
4
3-
 thermometers
 pH meter (or pH analysis kits)
)
 waste collection bottles for chemical analyses

D-ring plankton nets (1 per group)
 brushes for nets
 small nets for collecting material in buckets
 buckets (1 per group)
 labeling tape and markers
 sample vials (2 per group)
 ethanol
 sterile bottles for fecal coliform bacteria sampling (2 per group)
1.
Record the sampling location, date, time, and names in your group. We will sample in three areas with different ecological conditions (e.g., fastmoving, slow-moving, moderate flow), with 2-3 lab pairs in each area.
2.
Record general observations about the amount of vegetation covering the stream, how much light could reach the stream, color, turbidity (how much particulate matter), and qualitative flow rate (i.e., slow, moderate, fast).
3.
Measure the stream depth at three points across the stream, in a line that includes the location of your invertebrate sampler (see below) and runs perpendicular to streamflow. One of the three points should be at the location of the sampler.
4.
Water chemistry sampling :

Dissolved O
2
: In order to obtain an unbiased dissolved oxygen
(DO) sample, it is necessary to avoid air bubbles during the sampling and fixing process. Use the Hach DO water samplers to collect DO samples. Before you add the reagents for your DO analysis, make sure to practice taking and handling a DO sample.

Rinse water sampler with water from your sampling location before taking a sample. Each group should take two samples.
Samples can be analyzed in the field or transferred to a sample bottle and analyzed in the lab.


Use your water analysis kits to determine pH, dissolved O
2
(DO),
T, NO
3
-
, PO
4
3-
. Alternatively, a pH meter may be used to determine pH. Conduct each analysis twice and record data.
5.
Before you leave lab, exchange data with at least one other group who sampled in areas different from your own group, so that you have data for two areas with distinct ecological conditions .
6.
Macroinvertebrate sampling :

Sample aquatic invertebrates in areas where water is flowing (don't sample in still pools) and the streambed is rocky or has pebbles.
Make sure to collect two macroinvertebrate community samples from your sampling area.

Hold your collection net about 2 feet downstream from your feet and dislodge benthic invertebrates from pebbles by disrupting the stream bed with your feet. Dislodged invertebrates will then flow downstream into your collecting net. If you are sampling in a rocky area, lift rocks and scrape them to dislodge benthic organisms.

Next, partially fill a bucket with stream water, turn your net insideout over your bucket, and vigorously shake your net in the bucket water in order to dislodge invertebrates from your collection net.
Use a brush/spray bottle to remove any organisms that remain on your net after shaking.
 Remove and inspect any large items one piece at a time (I.e., twigs, leaves, pebbles, etc). Try to dislodge any clinging organisms back into the bucket before you discard the item.

Allow your sample to settle.

After your sample has settled, gently remove water from the bucket using a sample vial, in order to reduce the volume of water.

If necessary, use a second bucket to further 'concentrate' your sample into a smaller volume of water. Using the D-ring net as a filter, pour the reduced volume of water with invertebrate sample from the first bucket through the net into the second bucket. This operation can be repeated as many times as necessary, using increasingly smaller volumes of water, until your sample volume occupies approximately half of the sample vial.

Finish filling the sample vial with ethanol. This will preserve the invertebrates until the next lab when we will identify them. Be sure to label each vial with sampling location and your group name.
7.
Fecal coliform samples (Homework 3) : carefully submerge a sterile 1 L
Nalgene bottle at your sampling site, rinse the bottle with sample water, and then collect your sample. Label with the date, your group name, the

site, and replicate number. Store in the cooler for transport back to the lab.
 labeling tape and markers
 small sample vials
 brushes
 featherweight forceps
 pans
 dissecting microscopes
 invertebrate identification guides
1.
Pour the contents of your preserved sample vial into a pan.
2.
Look for animals, some of which are likely to be very small. Using the keys provided, identify aquatic insects to order ; for any other organisms, identify them as specifically as you can. If necessary, use the dissecting microscopes to examine the organisms.
3.
Record the number of individuals of each taxon you find in your lab notebook.
4.
Remove the animals with the soft forceps and place them in sample vials. Try to put similar animals into the same bottle.
5.
Make sure to account for 100 organisms from each sample.
6.
Exchange data with the same group(s) you obtained water chemistry data from
(if each group has two samples, you will have data from at least a total of four samples).
You will be writing a lab report that encompasses field sampling, water chemistry, and macroinvertebrate community analysis. The only analysis that we will not include is fecal coliform bacteria testing (which will be the subject of Homework Assignment 3).
In your report, as usual, you should include title, abstract, introduction, methods, results, discussion, and references (literature cited) sections. In addition to presenting your data , questions that you should address include:
Nutrients and abiotic factors: o Describe each measurement made, the results found (graphs may be helpful, especially when comparing the different areas sampled). Describe how these parameters can be used to draw inferences about watershed processes and water quality.

Macroinvertebrate community analysis: o Why is macroinvertebrate community analysis useful in assessing water quality? o Calculate the species richness and Simpson’s diversity index and plot rank-abundance curves for communities from three areas sampled. o Use species diversity indices and graphs to describe and compare macroinvertebrate community composition at different sites. Which sites had higher diversity? Why? o Did you find indicator organisms? If so, which ones and what types of ecological conditions do their presences indicate?