SUBSAMPLING AND
TAXONOMIC PROCEDURES EMPLOYED BY
STREAMKEEPERS OF
(revision of
Arthur J.
Frost
INTRODUCTION
This document attempts to show the subsampling and taxonomic procedures
that are used in dealing with benthic macroinvertebrate samples collected by
Streamkeepers of Clallam County.
Some of the procedures given are tentative (not yet subjected to review
by others); all may be considered subject to change (when such changes have
occurred over the course of Streamkeepers’ existence I have noted them; note also the revision date
above). The sources of
non-tentative procedures will be given immediately following the discussions of
them; most of these sources will be electronic mail documents received by
Streamkeepers.
It amazes me that a well-established taxonomic laboratory has not already
prepared a document such as this for general distribution (granted, I have been
doing this sort of thing professionally for about ten years but I do not
consider myself to be well-established).
The hope is that I am not “reinventing the wheel” by preparing this
document.
Copies of published work which answer questions raised below (or which
tell me how much “wheel reinventing” I am doing) would be
appreciated.
SUBSAMPLING
The main purpose of subsampling is to keep specimen counts to a
reasonable number;
Most of the subsampling that
When subsampling, the first task is to let the floating
macroinvertebrates sink; this is accomplished by keeping the sample in water for
a couple of hours before doing anything more to it. Once this is done, the sample may be
spread out onto the Caton grid, attempting to distribute the sample solids as
evenly as possible. For small
samples, a portion of the area may be used; a sample that does not take up at
least five grid squares should be subsampled in some other manner. A sample that contains sand or gravel
(this should not be the case for Streamkeepers, but it does happen) should have such materials
removed: evenly distributing sand and gravel on the grid consumes more time than
the subsampling would save, and cleaning sand out of the grid afterwards can be
particularly difficult. Sand and
gravel (if present) would be examined, and any macroinvertebrates found would be
retained for possible inclusion in the additional taxon search (which is more
conmpletely described below).
Once the organic portion of the sample is spread out on however much of
the Caton grid is needed, grid squares are selected in some random manner (I
usually use polyhedral dice: a twenty-sided die for the “down” coordinate, a
six-sided die for the “across” coordinate; a result that points to an unoccupied
or already-used square is ignored.
This procedure is best carried out well ahead of time, with resulting
grid[s] used as needed). The entire solid contents of a selected
square are removed from the grid in the following manner: the “cookie cutter” is
placed in the grid square to outline its edges; the scoop is placed along one
edge of the cookie cutter and the contents are routed to the scoop using a
paintbrush or some other tool (I prefer to use a narrow plastic scraper;
cleaning out a paintbrush so as to not miss anything is not exactly easy). The contents of the scoop are then
transferred to a petri dish or other container (I use a “sorting dish” made from
two PVC flat end caps: a gap is cut into each cap that is about as wide as my
field of view at 10X, and the two caps glued together at the gap) and sorted
using a binocular stereoscope.
Nematode and annelid worms are counted and recorded as they are sorted
(specimens of these taxa often break; fragments including one natural end point
count as one-half, intact specimens count as one); other taxa are simply counted
(the full identification phase can wait).
Terrestrial and semi-aquatic
macroinvertebrates are not included in
the count. A running count
is maintained as each square taken is dealt with; the count target is 500 (a
count up to 50 over the target is acceptable). Each square taken is completely dealt
with (final counts will almost always be some figure over 500); at least two
squares must be taken (it is not
impossible for a single grid square to hold 500 or more
specimens).
In the event that a specimen or
large piece of debris overlaps two grid squares. Large pieces of debris should be cut
along the grid square border once that is determined. Specimens should be shifted such that
they completely occupy whichever grid square initially contained the head
(assuming that can be determined); if that is impossible specimens may be
cut. If a specimen is cut, only the head is counted (for
nematodes and annelids see the procedure given above).
The number of grid squares actually used and the number of those that
were taken are recorded; these numbers are used to determine the percentage of
the sample actually analyzed with the following equation:
(grid squares taken/grid
squares used) X 100
There is a field in the
Streamkeepers database that accepts this information; the information is used to
estimate the total number of organisms in the sample.
It is possible for subsampling to fail (the penultimate square
undershooting the target, the last square overshooting the target by more than
50); if this happens, either the sorted organisms are placed in a smaller tray
(such as that described below) and a subsample taken from that, or all sorted
macroinvertebrates are identified and an electronic subsample generated (see
further comments below). In either
case, it must be remembered that the final percentage is in fact a product of
two initial percentages (either two iterations of the formula above, or one run
of the above formula plus a run of (specimens identified/specimens subsampled)
X 100).
Once the count target has been reached the non-worms of the subsample
would be identified and tallied, with the results recorded and the specimens
stored as usual. The used grid
squares that were not taken are then subjected to an additional taxon search. The standard for this type of search is
to examine what remains in the grid using some magnification for a set period of
time, removing one specimen of any taxon not found in the subsample. I consider this method all too likely to
miss taxa, and so the procedure I use for this search varies from the standard
(and is sometimes modified under exceptional circumstances). My usual procedure is as
follows:
1) Clean the grid, saving
all debris still therein, preserving the debris in 90+% alcohol; label this
container as apopropriate.
2) Wait 48 hours for the
remaining specimens to reabsorb alcohol (don’t waste this time; go on to the next
sample), then re-float the sample
remainder.
3) Examine the float under
magnification, extracting one specimen of each taxon not found in the
subsample. To make the task of a
Quality Assurance tech a little easier, these specimens are stored separately
(labelling the container with the site, date, and the phrase “Uncounted Sorted
Specimens”).
4) Examine the remaining
organic debris, extracting one specimen of each taxon not found in either the
subsample or the float discussed above.
I usually move specimens still in this debris that represent taxa that
had been found in a previous step into the “Uncounted Sorted Specimens” container;
this presents a QA tech with organic debris that is free or nearly free of
specimens (thus reducing the amount of time such a tech would need to look
through the debris). The one
exception to the preceding sentence involves a sample that is largely matted
filamentous algae; removing specimens from such is never easy, and so is done
only when required.
5) If sand or gravel were
in the initial sample, such would be sorted at this time, extracting one
specimen of any taxon not found in a previous step (the remaining specimens
would be referred to the “Uncounted
Sorted Specimens” container, for the same reasons given under 4) above).
This procedure does take
a fair amount of time (at least an hour after the initial wait), but I believe
that so doing has a better chance of finding most (if not all) of a sample’s
additional taxa than does the standard method described above. In
the event that a number of samples are in progress at the same time, adequate
labelling of all containers in use is essential!!!
When a Caton grid is too big for a sample that nonetheless is known to
hold more than 1000 specimens, an alternative method uses a small-area tray (we
have a transparent plexiglass tray with a grid area of eighteen square inches;
the underside of the tray is marked–and the markings protected by a sealant–for
the grid). This tray is small
enough to fit under a boom stand stereoscope; once the sample is evenly spread
out here some water would either be allowed to evaporate or would be siphoned
off (this to prevent drift as the tray is shifted under the stereoscope). Most of the remaining procedure as
outlined above would be followed (the dice used to determine selection of grid
squares are in this case two six-sided dice of different colors). The testing I have done of this method
suggests that it is best to use a 24-cell culture tray as an intermediate
destination, completely emptying the gridded tray (this helps prevent specimen
dessication).
If a sample after being completely sorted and identified is found to hold
more than 550 specimens, a subsampling computer program is used (one such may be
found at www.cnr.usu.edu/wmc;
My handwritten bench sheets for each reach all follow nearly the same
format, whether or not subsampling is done; one clue that subsampling has been
done to a sample is the letter “A” rather than a number entered for a given
taxon (indicating that taxon was found in the additional taxon search rather
than the subsample).
The Streamkeepers database records the taxa (and counts) found in the
subsample and the results of the additional taxon search in separate
fields; the first is numeric, the second boolean.
“TAXONOMY”
As can be seen from the heading, I consider the word “taxonomy” to be
misused for what I am about to describe; a better word would be
“identification.” I would restrict
the word “taxonomy” primarily to what Stribling, Moulton & Lester (2003)
called “research taxonomy” and that part of “production taxonomy” that is
concerned with preparing identification keys; the remainder of “production
taxonomy” would be called “identification.” That being said, the remainder of this
discussion will focus on the following: the kinds of organisms to be counted,
and circumstances under which an otherwise eligible organism would not be counted.
The samples as collected include not only benthic macroinvertebrates, but
terrestrial organisms that for some reason had fallen in, semiaquatic
macroinvertebrates, and fish; of these only the benthic macroinvertebrates are
to be counted. Streamkeepers
applies macroinvertebrate identifications and counts to a ten-metric Benthic Index of Biotic Integrity. With this tool the taxonomic rank
considered the final identification varies by group, as
follows:
Nematoda. Stops at phylum level. A
parasitic nematode must be completely free from its host to be
counted. A damaged specimen
must have at least one natural end point (those with only one are counted as
one-half).
Cnidaria, Platyhelminthes,
Annelida. Stops at class
level. For budding Hydrozoa, clumps count as one. Damaged Turbellaria are counted only if the
fragment includes the head (the usual standard) or the pharynx (the standard I
use); counts may be different depending on the standard used, which therefore
must be specified by the initial identifier. Damaged Annelida are treated in the same way
as damaged nematodes (see above).
Mollusca. Stops at family level. N.B.: originally it was believed that
Gastropoda were not taken that
deeply; this has resulted in some entries for “Gastropoda 1” in the Streamkeepers
database. Empty shells are not counted.
Arthropoda general
comment. Cast-off exoskeletons are not counted;
damaged specimens must include the animal’s head.
Arthropoda other than
Insects. Generally stops at
order level. For some subgroups of
the subphylum Crustacea, the “order
level” is not necessarily that currently accepted (for example, Copepoda, Ostracoda and Cladocera remain undivided; specialists
consider each of these to hold more than one order. A further note on Cladocera and Copepoda: at least one Streamkeepers
correspondent states that these should not be considered; as ecological variable
values are available, we are continuing consideration for the moment; we need to
get more opinions on this subject).
Insects. Generally stops at genus level. Exceptions are as follows: Plecoptera families Capniidae and Leuctridae; Coleoptera family Dytiscidae; Diptera families Chironomidae, Syrphidae, Sciomyzidae, Ephydridae (?) and Muscidae; all of these stop at family
level. For taxa with a pupal stage: pupae
(which are generally identifiable only to the family leve1) are included in the count only if one of the following conditions
is met: a) the taxon is
customarily identified to the family level; b) larvae identifiable to genus are not
also present in the sample; or c)
all larvae in a given sample are identified as the same genus (in which case the
pupae are assumed to also represent this genus). Condition c) may be deleted in the
future.
LITERATURE
CITED
Stribling, James B.,
Stephen R. Moulton II, and Gary T. Lester. 2003. Determining the quality of taxonomic
data. Journal of the North American Benthological
Society, 22(4):621-631.
December.