Crab Team Monitor Toolbox

Please submit data gathered each month electronically (as below) within a week of completing sampling. After your final sampling for the season, please mail us the hard copes of the data sheets in the brown, pre-addressed envelope you received them in at the beginning of the season. In case that envelope is no longer available, use the general WSG mailing address:

Crab Team
Washington Sea Grant
3716 Brooklyn Avenue N.E.
Seattle, WA 98105-6716

To submit data electronically each month:

1. Verify completeness of data sheets. Check that site metadata and volunteer hours are all completed. Ensure each row in the trapping data sheet is clearly totaled in the correct column. If you have questions about species identification, please get these answered before you submit your data sheets, so the data submission is fully complete.
2. Scan or take a photograph of data sheets. Your phone is often adequate for this, but quality can often be improved (and file size decreased) by using a scanner, or a scanning app on your phone (like Scanner For Me). Open all the photographs before sending
3. Rename files according to the convention. This might require transferring files to your computer, as not all phones allow you to rename files. All files should follow the basic format:
   • Site Number
   • Year (two digit, e.g. 17)
   • Month (single digit, e.g. 6, rather than 06 for June)
   • Content, either “Trap”, “Transect”, or “Molt” for data sheets, and trap type and number (corresponding to photo card) for trap photos.
   • Examples: 333.17.6.Trap.pdf  333.17.6.Minnow.1.jpg
4. Email all images to crabteam@uw.edu. Multiple emails are fine if necessary. File sharing websites like Google Drive and Dropbox also work for us. Make sure that if you have the chance to select the file size for photographs, that you choose at least “Large”. Selecting Medium or Small will result in pictures that are too low resolution to see details necessary for species identification.

We try to provide you with everything we think you will need at the beginning of the trapping season. Sometimes, however, you just run out of something, or data sheets are destroyed by the mud. We will do our best to replace any gear you have run out of. Just try to give us enough time to get it to you. If there is some piece of equipment that would make your monitoring easier, drop us a line, and we’ll see what we can do.

Our ID Guide (above) has many of the species that we think you are likely to see in your traps and molt surveys. However, every site will be different, and we can’t list absolutely everything. In addition, some groups, like sculpins, can be really hard to tell apart because there are many similar species. Part of our goal in this project is to have the highest quality dataset possible. With that in mind, we are always trying to identify every organism to the species level. If you find an organism, but aren’t sure what it is, contact us. We love playing Sherlock and trying to solve puzzles of mystery organisms! Take several photos from different angles and levels of magnification, and include some item for scale so we can see how big it is. If it is a molt that you can’t identify, you can actually keep the molt so we can get the specimen from you to be 100% positive.

On your data sheet, you can use a place holder, such as “Unknown species A/B/C/etc.” until we can figure out what the organism is. Then we can provide you with a species code to enter on your data sheet. Thus, don’t email us your data sheets until you have identified all the organisms to the species level. If you know the species, but it’s not on our sheet, you can still figure out the species code, remembering that the four letters come from the scientific name of the species, the first two letters of the Genus name and the first two letters of the species name: CAMA is CArcinus MAenas. But it’s probably good to write out the entire name for us as well.

Haphazard sampling, which is distinct from “random” sampling in the strict sense, is a common feature of many ecological studies. It is a practice that attempts to balance statistical rigor with field practicality, and reduce both human and crab bias in determining which crabs get measured. Reducing these biases is the best way to be sure that the subsample of crabs we measure closely matches the true sizes of all the crabs in each trap.

The short answer about the best way to do it is:

1. Close your eyes (reduce human bias),
2. Gently mix the crabs in the bin (reduce crab bias),
3. Select the first one that you grab and measure it.

Caution: You might want to modify this if you are dealing with larger, “pinchier” crabs!

For a (much more than you ever wanted) detailed explanation of why we do it this way, read the Protocol in Focus from the Crab Times Winter 2016 Volunteer Edition.

In general, females have wide abdominal flaps for carrying eggs that distinguish them from males. But in some cases it’s harder to tell than others. One particularly confusing scenario occurs with hairy shore crabs (Hemigrapsus oregonensis, HEOR) which can be infected with a parasitic isopod that feminizes the males – that is, it causes the males to look more like females in the shape of their abdominal flaps – read all about it in our Protocol in Focus piece on this parasite. Depending on the crab species, distinguishing between the abdomen flap size can be tricky, so we’ve created this handy Crabdomen Guide and a Protocol in Focus article all about distinguishing between crab sexes.

Wider than a typical male, narrower than a typical female. This crab is a male infected with a parasitic isopod that feminizes the crab. Record as if it were a male, but note in the comments section.

If you come across a HEOR that doesn’t quite look like either male or female, assume it is a “feminized male”, and count it as a male.  Record the carapace with (if it’s one of the 10 you haphazardly select to measure) in one of the 10 boxes, and include it in the total number of males for that trap. Though not required, you’re welcome to make notes of any feminized males you catch in the notes section for that trap.

As always, if you aren’t sure what you are looking at, take good notes and send us several close up photos to ID for you. Please send us these photos and questions before you submit your data, so the data sheet you submit is complete.

Though it may seem like an easy distinction, sometimes it can be tricky to discern whether the shell in your hand is a molt or a dead crab. To help you get to the bottom of it, we’ve outlined four questions you can ask yourself – Find more details about what to look for in this Protocol in Focus piece .

1. Is there any obvious trauma? Some predators will leave specific clues behind after munching on crab shells. Looking for these clues may help you distinguish between a molt or a dead crab that met an unfortunate demise.
2. How heavy is it? Molts are typically much lighter than dead crabs with tissue still in-tact.
3. Does it open? Because molting crabs essentially climb out the back of their shells, the shell of a molt should easily open on a hinge (much like a jewelry box). If it’s hard to open, it’s likely a dead crab.
4. What’s inside? Molted shells should be quite clean inside, while dead crabs may contain goo or crust of internal tissue. Remember that feathery gill tissue appears in molts too.

A molt (left) will have a clean shell interior, and open like a jewelry box hinging at the eyes. A dead crab (right) may be filled with “goo” of various colors in the middle section of the shell and attached to the inside of the carapace. Note that both crabs appear to have gills, while the molt actually only has the molted coverings of the gills. Photos Jeff Adams and Emily Grason

If you come across a shell that you’re not sure about, take some notes on your molt data sheet and snap some close up photos from different angles. Send them our way, and we’ll help you make the call. Please remember to send us these photos and questions before you send us your data, so that your data are complete upon submission.

The minnow and Fukui traps target different sizes and shapes of organism. The minnow trap has a smaller opening and mesh size, meaning it catches smaller critters than the Fukui. We use both traps because we are trying to get a snapshot of everything that lives in pocket estuaries. “Zeros” are also very important data points in our sampling scheme. So, even if we come up with empty Fukui traps, we are learning something: that there are no large crabs visiting that site. If we only used minnow traps, we would never be able to tell whether or not large crabs were present.

For a more detailed explanation, check out our blog post on this topic.

You might have noticed a green mat of vegetative material living directly atop the mud or on the stems of pickleweed, and been unable to decide whether it was wrack and washed in, or whether it was actually rooted in place and living there. This is a category of seaweeds known as filamentous green algae. We started tracking them in 2016 becauase they can grow quickly in response to high nutrient levels in pocket estuaries. The nutrients might be either part of the natural seasonal changes, particularly in spring, or they might come from nearby human activities. This is part of our program’s goal to track the health of pocket estuaries.

A blob (technical term) of filamentous green algae with hand for scale.

How do I identify filamentous green algae? Filamentous green algae is made up of very skinny individual filaments, and takes on a wooly appearance when growing on the mud. The color varies from dark conifer green, to yellow and white, as it bleaches when stressed by too much sun, heat, or desiccation. It can be partially covered with mud making it look somewhat brown, too.

As always, if you are not sure what you are looking at, whether it is filamentous green or something else – send us a photo, and we’ll help you sort it out.

This quadrat has filamentous green algae lying on top of the mud like a mat, coloring it green. You can still see some bare mud patches as well.

Where do I find this type of algae? This is not found at all sites, but it particularly thrives living at high tidal elevations directly on very fine sediments, i.e. mud. We see it most often very loosely attached to the mud between pickleweed stems, but also growing attached to the stems and branches of the rooted vegetation itself.

How do I record it? Filamentous green algae is most often recorded as step 2b. It can range from 0 – 100% independent of the other categories of cover in step 2 (rooted vegetation, live epifauna, and bare space). The other categories in step 2 apart from filamentous green must total 100% exactly, regardless of the amount of filamentous green.

The exception to this case is if the filamentous green is clearly part of a mat of washed up algae, and isn’t attached to either the plants or the mud. This can be hard to tell, because in many cases, it’s only very loosely attached at best. In most cases it will be attached, but if it is not, it gets recorded in step 1 as wrack – seaweed.

Filamentous green can range from 0-100 and is counted independent of the other three categories in step 2, which must total 100.

Where do I record it? As part of step 2b on the new data sheet downloadable in the list of supplies above.

Why is it separate from the other categories of cover? We realized it didn’t make sense for this category to be mutually exclusive from the others, because it measures a different quality of the habitat. The category of rooted vegetation estimates how much of the substrate is stabilized by roots of living plants, or how prone the area is to erosion – something green crab could change. Filamentous green algae doesn’t contribute to sediment stability because it’s so loosely attached. It’s really more about how much light and nutrients are available. If we were to include it in the other categories of cover, we might be overestimating the stability of the bank at a site where filamentous green is abundant.

Limpets, mussels, and barnacles all count as epifauna

In 2016, we added the category of “Live Epifauna” to Step 2 of the quadrat estimation. We ask what percent of the space is taken up by live epifauna, which means animals living on or attached to the surface. This could be mussels, barnacles, snails, limpets, chitons; our rule of thumb is if it can’t move fast enough to leave the quadrat while you are observing it, it gets counted as live epifauna.

For more information on how to distinguish live epifauna from dead epifauna, and what we can do with these observations, read our Protocol in Focus on the topic.

It can appear that the shoreline doesn’t change much over the course of six months, so we are sometimes asked whether maybe only surveying the shoreline once per year is sufficient. We do the survey monthly because, just as on land, conditions in the intertidal environment change throughout the season, such as light and nutrient availability, water temperature, energy, and chemistry. All of these factors can cause slight changes that, on the scale of a single site from month to month, aren’t readily perceptible. But when we aggregate them over space and time, we can start to see patterns that aren’t obvious to the naked eye.

Check out what we learned in 2016 from our transect surveys in our Protocol in Focus on the topic.

Crab Team monitors certainly aren’t afraid of getting muddy, and estimating the roots of vegetation – instead of just the canopy – during the transect survey can be one of the muddiest tasks each month! So why do we care about how much of each quadrat has rooted vegetation in it? Read this Protocol in Focus blog post to find out.

Wondering what happens to all the valuable-as-gold information you collect each month after you click “send”? Read all about it in this Protocol in Focus blog post.

Believe it or not, quite a bit of thought went into what type of bait we would use for monthly Crab Team monitoring. Read this Protocol in Focus blog post to learn more about how we landed on mackerel as our bait of choice!

In addition to creating a scientifically robust sampling scheme, and one that is sensitive to finding green crab when they are rare, Crab Team also put a great deal of thought and guidance into the protocol with the goal of being good stewards of Washington’s shorelines. Even though our goal is to detect green crab as they first arrive and help avoid ecological damage, it is certainly true that the act of sampling itself can be a disturbance to a site. Repeated visits to muddy habitats can trample vegetation over time. Being caught in a trap can be stressful for the native crabs and fish. Multiply this by 56 sites, six times per year over successive years, and the cumulative effects, if we weren’t careful, could be substantial, and even reduce the resilience of sites to green crabs.

Read this Protocol in Focus blog post for our list of site stewardship best practices to be a great stewards of your site! Following all of these tips helps ensure that we can continue to monitor our Crab Team sites in the years to come.