Where and when do I submit data or data sheets?
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:
Washington Sea Grant
3716 Brooklyn Avenue N.E.
Seattle, WA 98105-6716
To submit data electronically each month:
- 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.
- 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
- 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
- Email all images to email@example.com. 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.
"How do I get more ______?"
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.
How do I decide what time to schedule monitoring?
A basic outline of the important things to think about when scheduling monitoring is included in the Volunteer Handbook (above). Shoreline surveys can be conducted any time that the tide is low enough to expose the habitat boundary and you will need to think about the tide cycles for trapping surveys. In either case, the key piece of information is knowing the tidal elevation your site. In particular consider the height of your habitat boundary and the height of your traps. Then looking at a tide chart will help you plan. A good website for tide charts is NOAA Tides and Currents, and TideGraph is an excellent app for iOS. Any time that the tide is predicted to be lower than either your habitat boundary or your traps, they will be exposed and you can monitor.
Ideally, you want to make sure the traps are submerged in water for the entire time they are set, so that (1) they are always fishing, and (2) any live organisms are constantly kept in water and will be in good condition to be released. So, you will need to first check that the second low tide of the day won’t drop low enough to strand your traps high and dry and unattended, which could provide an opportunity for raccoons and opossums to find them and try to go fishing themselves. You’ll also need to plan to return to the site in time to process all the trap contents as the tide drops. This planning is easier if your site retains some water on even the lowest tides, like a lagoon or a deep channel, and we’ve tried to select sites where this is the case for that reason. If this is the case at your site, you will have more flexibility about when you can return. (Coming soon: Mini tutorial on timing your sampling)
How do I record a species I can't identify?
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.
What do you mean by selecting crabs "haphazardly"? Why does it matter?
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:
- Close your eyes (reduce human bias),
- Gently mix the crabs in the bin (reduce crab bias),
- 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.
Is this crab a male or a female?
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.
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.
We do like to track these observations, so please include a tally of how many males were feminized in the comments column for that trap on your trapping data sheet.
As always, if you aren’t sure what you are looking at, take good notes and send us lots of 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.
Why do we use two types of trap?
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.
How do I record filamentous green algae?
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.
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.
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.
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.
What is "Live 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.
Why do we survey the habitat transect every month?
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.