Stream and Lake Monitoring Protocol

SOP 13 – Macroinvertebrate Processing and Identification, v. 1.0, Page 1 of 16

Stream and Lake Monitoring Protocol

ARCTIC Network (ARCN), Alaska

Standard Operating Procedure (SOP) # 13

Macroinvertebrate Processing and Identification

Version 1.0 (November, 2007)

I  Revision History Log:

Prev. Version # / Revision Date / Author / Changes Made / Reason for Change / New Version #

This SOP provides instructions for the laboratory analysis of macroinvertebrates collected for the stream and lake monitoring in the five National Park units in the Arctic Network (ARCN). These methods have been modified from ABR, Inc. (Cole 2002, 2003).

II  Sample Preparation and Identification

STREAMS

Sample Log In

  1. When samples arrive in the lab, immediately assign a sample number and enter number into the Sample Log (Figure 1). Log information to be entered at this time includes site name, sample point, collection date, and sample number. The sample number (1, 2, 3, 4,…) is assigned to each sample of a given project in the order in which the samples are entered in the log (they may be different from site numbers or codes).
  2. Use a sharpie to write the sample number on the outside of the bottle or whirl-pak. If the sample is stored in more than one bottle, use the same sample number, but write “(1 of 2)” or “(2 of 2)”, etc.
  3. When duplicate samples from the same field site are taken, they should receive different sample numbers, with the notation “(DUP 1)” or “(DUP 2)”, etc. following the sample number. This information, together with the site number or code, will provide the needed information to accurately identify each sample.
  4. Samples should be stored in appropriate location in lab in secondary containment until they are sorted.
  5. Before being stored, they should be checked to ensure that the proper volume and dilution of alcohol has been added. Sample bottles should not be more than 2/3 full of sample material, and should be well covered by alcohol that smells strong. A weak or lack of alcohol smell indicates that the alcohol is likely too diluted and that it should be replaced. To replace the alcohol, pour the sample into a sieve over alcohol waste container, and refill with 80% ethanol alcohol.

Sample Preparation

Samples from the field are often laden with fine materials that cloud the sample and very coarse materials that add a lot of volume to the amount of material to be sorted through. It is the aim of sample washing to rid the sample of these materials at the extreme ends of the size spectrum without losing any macroinvertebrates. Importantly, some samples require a lot of washing, others require none. Most require some; only experience will help determine in advance how much washing will be necessary.

If the sample contains lots of silt and sand (which can be seen through the bottle once it’s settled), much of this can be washed out through a 250-m sieve:

  1. Pour the sample contents onto a set of nested 1000-m and 250-µ sieve rinsing the sample container to ensure that all materials have been transferred onto the sieve.
  2. Run tap water through the sieve and retained material for about one minute; use water stream to concentrate sample in one corner of the screen.
  3. Dump the sample into an examination tray, rinsing all material retained on the screen into the tray.
  4. Fill the tray half way with water; allow the water to stir up the contents of the sample.
  5. Swirl the tray gently and pour again into the sieve, this time allowing the water to run into the sink, while catching the sample in the sieve.
  6. Repeat this process until the water from the sample is clear.

At this point, the sample may have large material that can be hand picked, rinsed into the screen and saved for benthic organic matter analysis.

  1. Pour sample again into the tray and fill halfway with water.
  2. Hand pick large leaves, sticks, etc. from the sample and save for benthic organic matter analysis.
  3. Closely examine, and wash (if necessary) items that have invertebrates adhered to them.

Finally, if the sample has a large amount of gravel and coarse sand, this material can also be separated from that to be sorted:

  1. Again fill the examination tray halfway with water and slowly pouring water and the sample into the sieve with gentle swirling action (Figure 2). The idea is to pour water from the tray at a rate that carries the invertebrates and other lighter organic material into the sieve, while heavier pebbles and sand are retained in the tray.
  2. Repeat this process, leaving the sample in the sieve a number of times until the water isn’t carrying any smaller inorganic materials out of the tray when you are pouring.
  3. This process may sometimes require pouring out and refilling the tray with water 5 to 10 times.

Once the sample has been washed the pebbles and coarse sand should be thoroughly inspected for caddisfly cases and mollusk shells that are also retained in the tray (Figure 3). If these are seen, then the same proportion of this material will have to be searched as was searched of the washed sample. Save this material and put whatever wasn’t searched with a dissecting scope back into the “unsorted fraction” bottle. At this point the sample will be separated into 2 fractions. All material greater than 1000-µ is the coarse fraction. All material >250-µ but <1000-µ is the fine fraction.

Sorting Procedures

  1. All material from the coarse fraction should be examined under a dissecting microscope under 10x (minumum power). Organisms should be identified to the lowest practical level, usually genus.
  2. All material from the fine fraction should be examined under a dissecting microscope under 10x (minumum power). Organisms should be identified to the lowest practical level, usually genus. If there is a large amount of fine material and many organisms, the fraction can be subsampled using a folsom splitter or gridded petri dish (see step 3).
  3. Randomly select one square and transfer into a Petri dish. Add water to the sample, just enough to facilitate sorting through the material, but not enough to spill easily. Usually, about half full is enough (Figure 5).
  4. Place the Petri under a dissecting microscope. 10x power is the preferred magnification for sorting.
  5. As a rule, if the entire depth of the material is not in focus, then there is too much material and/or liquid on the Petri dish. In this case, carefully place half of the sample on another Petri dish and sort after the first half is sorted. This is important in ensuring that the sample can be well searched and that organisms aren’t being missed because they are out of focus or because too much material is impeding visibility.
  6. Use a gridded Petri dish or a Petri dish with four dividing ridges to provide reference points while the sample is being searched. Beginning on one side of the dish, begin systematically working through the material within each grid piece or section, until the dish has been entirely searched. This systematic approach ensures that the entire sample is searched, and searched with equal effort.
  7. Once the entire Petri dish has been searched, scan the dish for another 30 seconds or so to ensure that organisms were not being missed. Look for small organisms, including nematodes, microcrustaceans, and midge larvae that are more easily overlooked.

8.  Once finished, the sorted residue should be placed in a tin weighing boat and dried at 60°C for a minimum of 48 hrs for later use in benthic organic matter estimates.

9.  DESIRED REMOVAL RATE: 95% (i.e. only 1 out of every 20 invertebrates should be missed, preferably fewer)

Rules

Samples will primarily contain aquatic macroinvertebrates, but will also contain:

• terrestrial invertebrates (spiders, commonly)

• airborne adult stages of aquatic invertebrates

ALWAYS REJECT: Do not sort or count these organisms – i.e., leave them in the sorted residue.

Larval Molt Skins: Look for the split down the ecdysial line on the mid-dorsum of the specimen. Skin casts are usually transparent/translucent and do not have any internal materials.

Invertebrates Obviously Dead before being preserved.

Eggs of invertebrates or vertebrates (Oligochaeta worm egg cases are not counted)

Miscellaneous body parts: legs, antennae, gills, tips of abdomens, etc. Counts are based on heads and thoraxes.

Small fragments of Oligochaeta: Worm counts are based tips, when worms are fragmented (more often than not). Count only large pieces, and tips (anterior and posterior ends), count each tip as one half. Ignore smaller sections that do not have an end on them.

SORT, BUT DON’T COUNT: Put these specimens in a separate vial for a taxonomist to see. If large numbers of any of these are encountered, but a number of them (6-12) in a vial and make a note estimating the total number you encountered.

Empty caddisfly cases

Empty mollusk shells

Vertebrates: Fish and amphibians (including tadpoles) should be saved, and counted separately from invertebrates.

Headless specimens should be sorted, but not counted.

Taxa Expected to be Regularly Encountered in the ARCN Network
Sci Name / Common Name / Notes
Oligochaeta / aquatic worms / count large portions of bodies as 1, tips as 1/2
Chironomidae / Midges / look for pupae, as well
Nematoda / Nematodes / look out - these are easy to miss!
Copepoda / copopods / can be very small
Daphnia / water fleas / Can be very small
Ostracoda / seed shrimp / Can be very small - look like tiny white jelly beans
Odonata / dragon/damselflies
Trichoptera / Caddisflies / sort, but don't count empty cases
Molluska / snails/clams / sort, but don't count empty shells
Hemiptera / true bugs / see sorting notes for water striders, sort and count Corixidae
Coleoptera / Beetles / sort and count all beetles and their larvae
This list is almost certainly incomplete - look out for other organisms, as well

Remember: Counts are based on heads – sort otherwise intact bodies, but don’t count them.

Use of Sorting Vials

Invertebrates should be sorted into 1-dram snap cap vials. Use separate snap cap vial for each distinct group of organisms. Sometimes, larger containers will have to be used for mollusks and other larger organisms. Generally, 6-7 vials are used per sample, sorted into the following groups:

• mayflies, stoneflies, caddisflies, dipterans, beetles, mollusks, other.

Depending on the relative abundance of other organisms, additional vials may be used, or other groups substituted for one or more of those listed above. Importantly, place a small strip label with the LAB SAMPLE NUMBER inside of each vial. This way, if the vials ever become separated, they can still be identified.

The SORTED INVERTEBRATE VIALS should be bound with a rubber band (include both vials, coarse and fine fraction and any extra; affix a SORTED SAMPLE label to the sample vials (Figure 6).

To recap, these products are expected from each sample, unless otherwise instructed:

• Vials with sorted invertebrates (include label)

• Vial with large/rare organisms (include label)

• • Sorted residues (may sometimes discard, check with project manager)

• COMPLETED INFORMATION IN SAMPLE LOG

LAKES

Use sugar-buffered Lugol’s solution (3 ml per 50 ml of sample) to preserve zooplanktons samples. Use formaldehyde (5 ml per 50 ml of sample) or 95% Ethanol (20 ml per 50 ml of sample) to preserve benthic invertebrate samples. Zooplankton and benthic invertebrate samples are processed in different ways:

I. Identification and Enumeration of Zooplankton and Cladoceran Eggs

Recipe for Lugol’s Solution (sugar buffered)

Using a funnel, add 10g iodine, 20g potassium idodine, 200 ml distilled water, 20 ml glacial acetic acid, and 130 g sugar to a 250 ml light-proof nalgene container. Tighten lid, shake to mix and allow all sugar to dissolve (note, this may take several hours). Use this solution as a stock solution and take a smaller volume in the field. Note, solution should be saturated with sugar

1. Note on data sheet:

· Lake, station, and depth sampled

· Date sampled and sampler

· Net used (diameter of net opening)

· Date counted and counter

· Scope and magnification used

Pour the sample into a plastic cup fitted with a 70um nitex screen. Use a wash bottle to rinse the sample jar making sure all zooplankton are rinsed into the screened contained. Use the wash bottle to rinse as much of the preservative out of the sample by running water through the sample and the screened bottom of the container. After the sample has been washed, use the wash bottle to put the sampled zooplankton into a plastic counting tray (approximately 8 cm X 8 cm X 1 cm deep).

2. Count all individuals of each taxon in the sample. For taxa with more than 100 individuals in the sample, sub-sample as follows:

· Dilute sample to 100mL in a graduated sample jar.

· Be sure to homogenize the sample before sub-sampling (stir it up).

· Use a 5mL Henson-Stempler pipette to obtain a 5mL/100mL sub-sample.

· At least 30 individuals (pref. 50) of each taxon you are counting must be present in the sub-sample (600-1000 indv. / full sample).

· For borderline sub-samples, you may subsample twice and either: 1) add them for a 10mL/100mL sub-sample (be sure you do NOT dilute again before taking the second sub-sample in order to maintain the same concentration); or 2) take the mean of two sub-samples in which case you will count one sub-sample, then remix the entire sample, and then sub-sample and count again.