Frequently Asked Questions
The Answers to Some Common Queries
Did something related to Biologica’s work put you in an inquisitive mood? Check here to see if your question has been answered before, and if it hasn’t, we would be happy to answer over social media or e-mail. If you have found a mysterious critter that you would like to have identified, or just have a question, please post it to the Biologica Facebook page, Tweet it to @biologicabc, or post it to Instagram and tag @biologicabc.
I found this thing. What is it?
GREAT question. Perhaps our favourite question.
Step 1: Take a photo of it (remember that the better the quality of the photo, the better our chances of being able to successfully identify it).
Step 2: Post the photo to our Facebook, tag us in an Instagram post (@biologicabc), or mention us in the tweet and we’ll put our heads together to get you an answer.
What are my options for ID level and what are the differences among them?
For some projects, identifying the organisms in a sample to genus or family-level (less specific) is acceptable and because it is often less labour-intensive than identifying to species-level (more specific), this can be done at a lower cost. We encourage our clients to be sure that their chosen levels of identification effort will satisfy the scientific reporting needs of their projects before commencing analysis.
Can I perform taxonomic analysis myself?
Maybe! For larger-bodied taxa in particular (e.g. whales, vultures, predatory cats, ungulates, trees) field guides with taxonomic keys are available and can be used without any specialized equipment. For organisms possessing distinguishing features indiscernible to the naked eye, specialized training and equipment are likely required in which case you should contact us. There are also cases where identifying an organism can only be done using genetic markers, a process for which we may also be of assistance.
When can we expect our results?
Our standard turnaround time is 90 days from the arrival of samples at our facility. We are able to process samples more quickly at certain times of the year, and depending on the size of a project. Please contact us for an accurate delivery date.
How are our data reported, and can we request different formatting?
Our data is reported in excel spreadsheets. We include the data in both column format as well as a matrix format. We are able to reformat data to fit most of our client’s databases. Please contact us if you have any specific data requirements or requests and we would be happy to accommodate you.
What are Benthic invertebrates?
Click the links below for more information.
When you say “mesh size” or “screen size”, to what are you referring?
When benthos samples are prepared for analysis they are rinsed through screens to remove silt/dirt/sand/other sediment. The different screen sizes refer to the gaps in the screen (often 0.5 mm (AKA 500 microns) or 1.0 mm in marine sediments, and 250, 400, or 500 microns in freshwater sediments) through which organisms smaller than those gaps may pass.
Which mesh size should I use for my project?
This question will vary depending on factors including (but not limited to) sample substrate, types of organism in the sample, organisms being targeted for the purpose of the project, and analysis protocols (such as CABIN). A very important consideration is also what the mesh size has either been historically for the project, and/or what historical data may be used for comparison. If you would like guidance regarding field protocols, please contact us.
Where do we get screening apparatus?
We fabricate and sell screening equipment, so please feel free to contact us if you would like to purchase screening equipment. We also rent equipment and trained field staff if you would like to have us help with your field program and ensure the quality of your benthic samples.
What are the different samplers and why might I pick one over the other?
The most frequently used samplers for benthos collection are Van Veen grabs for marine work, and petite Ponar/Ekman grabs, Hess or Surber sampler, or kicknets for freshwater work. Grabs are typically deployed from a boat, and are used for samples collected from deep water or depositional environment. The different types of grabs collect different volumes of sediment. Hess, Surber, and Kicknet samplers are used in running waters (e.g. streams/creeks/small rivers), mostly in nearshore or shallow waters. Kicknets are used for surveys that utilize the CABIN protocol.
What would you expect to see in a benthic sample?
The biotic communities in marine, freshwater, and estuarine environments can vary highly with numerous factors including geography, sediment type and particle size distribution, water depth, water and sediment chemistry, temperature, predation, pollution, etc. Variability can be observed even among distinct stations within a single waterbody, or seasonally at a single station. For some examples of organisms that we find in benthic invertebrate samples, see our freshwater or marine benthos pages.
Can we have our samples processed without sorting? Can we do the sorting?
Sorting is the process of separating organisms from debris and as such, thorough and careful sorting is critical to ensuring data are of the highest quality. Microscopic organisms are often found attached to or contained within debris or organic material that may be mistakenly disposed of, an error which could potentially confer a reduction in species diversity, abundance, and biomass estimates. As a result, sorting should be performed carefully by trained technicians using microscopes, and subjected to stringent quality control measures.
What are phytoplankton?
Click here for more information.
How are phytoplankton collected?
Phytoplankton are collected within water samples, which can be collected using surface grabs, at discrete depths using specialized devices such as Kemmerer samplers or Niskin bottles, or within whole-column water samples.
How does one preserve phytoplankton?
Phytoplankton are preserved within samples using “Lugol’s Solution” (or “Acid Lugol’s” if it contains acetic acid). Lugol’s solution contains iodine which stains cells for identification. For more information on Lugol’s please contact us.
How much Lugol’s should I use?
Typically a 0.5 -1 mL preservative per 100 mL sample will be fine, although volume of preservative necessary for adequate preservation will vary with abundance of phytoplankton within the sample. Please refer to the photo on our phytoplankton page which presents a rough guide for the colour your sample should reach with the correct amount of Lugol’s.
What are zooplankton?
Click here for more information.
I would like to collect zooplankton, how should I do this?
Zooplankton are collected using nets, which can vary depending on the sampling environment and target taxa, but a good place to start is Dynamic Aqua in Vancouver. Although we don’t currently fabricate nets, we can most certainly offer guidance on your sampling protocols, so please feel free to contact us.
Why do people put alka-seltzer in zooplankton samples?
Alka-seltzer is used to anesthetize the zooplankton and help them retain their shape and minimize egg loss, which can aid in identification. We do not currently know the origins of this funny practice, but if you do, please contact us to let us know. Alternatively, clear carbonated beverages may be used including (but not limited to) club soda, tonic water, 7Up, Crystal Pepsi, or Clearly Canadian. We recommend this practice as we have found it to result in high-quality samples.
What is sugar formalin and why do people use it?
Early research on preservation of zooplankton noted that a sucrose-formalin solution could be used to reduce body-shape distortion and egg loss in cladoceran taxa. As such, it is not unusual for limnologists in particular to use sugar formalin for preservation of zooplankton samples. Some field programs still send us samples in sugar formalin, some in normal off-the-shelf 10% buffered formalin solution, and some in ethanol. If maximizing cladoceran taxonomic diversity or reducing egg loss are priorities for your zooplankton sampling program, it would be worth learning more about the different preservation practices and how they might work for your program. Alternatively, contact us.
How do we get these fish stomachs to you?
Following removal from the fish, stomachs can be shipped to us in preservative (10% formalin is recommended) without refrigeration.
What sample containers do we use?
We prefer leak-proof containers (jars/vials/sample cups). Whirlpaks may be used if these are stored in a large leak proof container (inside the shipping container) such as a bucket with a lid or a double-bagged Ziploc.
No-one here is able to remove the stomachs. Can we send you whole fish?
Absolutely. In this case, you will need to freeze or preserve the fish whole. Please note that we have limited freezer space for large-bodied fish, so please contact us if you have a large number of specimens are >15cm and you do not intend to remove the stomachs. There will be a slightly greater cost for analysis given the increase in labour times, and please bear in the mind the cost of shipping may be affected.
What is formalin and how is it different from formaldehyde?
Formalin is a solution of formaldehyde in water. Formaldehyde or “full-strength formalin” is only 37% formaldehyde in solution, so a 1:1 mixture of full strength formaldehyde to water, or, 50% formalin, is actually 18.5% formaldehyde. Most samples are preserved in a solution of 10% formalin, which is just under 4% formaldehyde in solution with water and methanol, often buffered with phosphate.
How important is it to buffer formalin, and what buffering agent should we use?
Non-buffered formalin can eat away at calcareous structures such as the shells of molluscs. These structures may be important for accurate taxonomic identification, thus their degradation may result in lower-quality data. If you don’t purchase pre-buffered formalin, it is easiest to use Borax (Sodium Tetraborate). You will likely need 1-2 Tbsp per sample to reach pH 7, depending on the acidity of your starting solution. The buffering capacity may decrease with time, so it is important, particularly in benthic samples containing molluscs, to transfer the sample to alcohol within 7-10 days for long-term storage or analysis. The addition of calcium carbonate chips (“marble chips”) can extend this storage time.
Do I need to use formalin, or is ethanol sufficient?
Formalin is a “fixative”, thus creates more robust and high-quality specimens than does ethanol, which is merely a preservative. For this reason, we highly recommend the use of formalin in samples that contain soft-bodied taxa (e.g. annelids, cnidarians in marine benthos or zooplankton) or samples that require pigmentation patterns for identification (e.g., ichthyoplankton). Samples containing primarily organisms with exoskeletons (e.g. freshwater benthos and zooplankton) may be preserved in ethanol with minimal effect on the data, particularly if there is no particular interest in soft-bodied taxa (e.g. oligochaetes in freshwater benthos). For information regarding the safe handling of preservatives, please contact us.
Can I/you ship preservatives to our field staff?
Possibly, depending on the destination and time we have to deliver. The Dangerous Goods status of the desired preservatives is the primary concern, as due to their flammability and toxicity certain preservatives will not be carried by some couriers and expediters. As a quick rule of thumb, 10% formalin is relatively easy to get onto a plane or a truck, whereas other preservatives can require more effort. Please feel free to contact us for more info.
Where should I send these samples?
See the contact us page for more info, but in general:
488-F John Street
We receive shipments from all major and minor couriers, and there is someone here to receive samples Monday-Friday 8:30-5:30 and weekends by special request. We can also pick up shipments from Greyhound and the Victoria International Airport (YYJ). Make sure you contact us prior to shipment if you are interested in having your samples picked up.
How should I pack my samples and what documentation should I include?
Please make sure jars are closed tightly, ensuring the threads of the jar are void of debris or dirt that would interfere with the seal of the lid. A few wraps of electrical tape around the closure is also a good safeguard against leakage. Ensure all sample jars are clearly labeled with a sample number, date and the number of jars per sample. Generally, samples preserved in ethanol or formalin do not need to be refrigerated, but they should be kept out of direct heat. We encourage our clients to include applicable MSDS documentation with the shipment. As part of your shipment, please include a packing inventory (whether as part of a COC or on its own), and any specific handling or analysis instructions that may not have been part of the quotation process. You may also contact us regarding handling and/or analysis.
Do we have to preserve our samples before we send them to you?
Unless samples can get to us in such time that we are able to preserve them in the lab without risk of degradation (and arrangements to this effect have been made in advance), then samples must be preserved in the field. If you are unsure, the answer is probably yes.
Do these samples need to be refrigerated?
Samples preserved in ethanol or formalin do not need to be refrigerated, but they should be kept cool.
Can we send you sediment samples for benthos analysis without screening them?
Only under specific circumstances (samples must be small, and collected on the same day). On rare occasions it may be feasible to send us unscreened samples that have been preserved, but this must be discussed with us beforehand. This practice isn’t encouraged as it is often more costly than performing screening in the field. In general, the quality of sample may also suffer. Please contact us to discuss your options, or, alternatively, please consider contacting us regarding having a Biologica staff member assist you in the field.
Video Benthic Analysis
How do you survey under water?
Dive video can be taken along transect lengths as well as photographs. Both of which can be analyzed for community composition.
How do I ensure I get the best identifications and analysis possible from dive videos?
Transects of similar length should be placed along the survey area. Water depth should be recorded at the start and end of the transect. Divers should attempt to maintain a slow constant speed, distance from the substrate, and position relative to the transect line throughout the dive survey. This will ensure the area analyzed is constant among the distance intervals, within the transect, as well as a more detailed analysis of the substrate composition and substrate slope.
How do I ensure I get the best identifications and analysis possible from dive photographs?
If possible use a tripod to ensure distance from the substrate is similar across all photographs. Adequate lighting is necessary so use of a flash is always recommended.
When is the best time to take dive videos?
Avoid sampling in low visibility conditions (at night or during algae blooms). Natural light is preferred but dive lights can also be used to aid in visibility.
There’s mould in my house, can you help?
We’re very sorry to hear that. However, no we can’t. Contact our friends at HazPro Environmental.