Plastic debris can come in all shapes and sizes, but those that are less than five millimeters in length (or about the size of a sesame seed) are called “microplastics.”

Following extraction, visual counting with an optical microscope is the most common technique for quantifying microplastics; a technique that is labour intensive and prone to human error. Spectroscopy (FTIR; Raman) are the most commonly applied techniques for identifying polymers collected through visual sorting.

if you want can see link below

https://ebrary.net/24934/environment/identification_microplastics

https://www.gammadata.se/assets/Uploads/Microplastic-and-Raman.pdf

https://bura.brunel.ac.uk/bitstream/2438/17133/1/FulltextThesis.pdf

You are correct, visual sorting gets increasingly difficult as the particle size gets smaller. The sizes of MPs that you are able to pick out of your sample first comes down to what you can see, and that is often dependent upon the magnification abilities of your microscope. And there can be a fair amount of error associated with that as MPs often look like other things (e.g., diatoms). Adding additional techniques before visualization can help a lot.

First is the digestion of the tissue. I have tried both KOH and H2O2 + heat on fish tissues and found them both to be effective. I typically use H2O2 at 65C for a few of hours with periodic agitation, depending on the size of the tissue sample. Karami et al. (2017) has a nice paper on different types of digestions. Next is separation from the surrounding media. If you are interested in separating by polymer type, then you can consider a density separation. Li et al. (2018) provides a good method. Just know that some of the chemicals used can be a little difficult to handle and particle size can impact buoyancy. The latter might be solvable by adding centrifugation (see Nguyen et al. 2019). You mention filtration, and I would say that is the most common method. There is some discussion about how to best filter samples to get the most MPs while avoiding contamination. While not the only one, Cai et al. (2020) addressed that subject recently. Personally, I think that filters are a good way to go if your MPs are large enough to be caught by it. You should consider passing the digestate and subsequent filtrates through multiple filters with smaller and small pore sizes so that you you don’t clog filter pores and when you get to the smallest particles large bits aren’t obscuring the view of smaller particles. Nanoplastics are still a big problem. The Nguyen et al. (2019) study says that their technique is able to separate those too, but I haven’t tried it yet. Its generally agreed upon (as of now) that there is no one good method to separate out the really small nanoplastics. And if you think you have a separation method, once they get that small, the only way to verify if you got any is by using an electron microscope (maybe uFTIR…very much maybe). That’s one of the reasons most people purchase fluorescent NPs to use in their exposure experiments. Next is the Nile red staining that you mentioned (I’m assuming you are using protocols from Maes et al. 2017 and Shim et al. 2016?). I certainly see this as one of the more commonly used methods to differentiate MPs from their background. And, if your microscope has enough resolution, you should be able to see particles

Melissa Chernick first of all, I'd like to give you a BIG thank you for taking all that time to explain all your suggestions, and even addressing the links to the papers you mentioned. Thank you.

And well, getting into the question. My main objective is to determine if there is or not MP's in shrimp, but since the work won't constitute a thesis or a paper (is more like an academic work that eventually will derivate in a short communication-like manuscript, not necessarily something very rigorous, but well made and peer-reviewed of course), I want to try something that doesn't involve the "musts" in MPs determination (FTIR, Raman, confocal or electron microscopy). These methods will be used, for sure, it is only matter of time and logistic to do a collaboration with another labs with the equipment, gathering students, technicians, researchers and experts in the field to do a more comprehensive analysis. But in this particular matter of my question I initially wanted to determine MPs only by visual sorting. Why in shrimp muscle? because it is the edible part of the shrimp, and a matter within food security. But I ran into this issue of the particle size. I, on the other hand, was thinking of trying with a bigger tissue, like stomach, to catch for bigger MPs that allows visual sorting. But I think that working with shrimps is going to end in recuperate MPs smaller than 500 microns. And there are papers that did visual sorting (with stereoscope) of MPs from abdominal muscle digestate, but they coupled that with spectroscopic methods and fluorescence microscopy (Fernández-Severini et al., 2020; Valencia-Castañeda et al., 2022). If I hypothetically still go for abdominal muscle, and catch MPs no less that 2 microns (I have filters of that pore size), do you think, in your experience, that will be possible to do a visual sorting classification? (size, color, shape, abundance). And in this context, is there a standardized procedure to perform visual-sorting classification?, meaning... I have the filters ready to be watch and... at which point one can stop counting, searching, looking for colors, etc.? would it be by drawing quadrants onto the filters and working a representative sample of each? Regarding fluorescence, you mentioned I should determine if I will get autofluorescence within the same wavelengths as the stain, sorry for my ignorance, but what does that means? I never used fluorescence microscopy before. Is possible though, to collect hemolymph from shrimps, to my knowledge, but in this particular case, it is not my main purpose. I will check on all those links you provided me, and the tip of using several pore size filters is a (very) possible route I will take on, in order to cope with clogging issues.

Thanks again for your answer, I guarantee you that it'll be of great help, as well as the documents and comments made by Ahmad Al Khraisat . That thesis work is very complete and it already provided me some answers to questions I didn't know I had, thank you so much to both of you.

Melissa Chernick sorry if I make to much questions, I know is part of the job to search for answers in science, but I'm very new in this field, and MPs involve lots of microscopy expertise I don't have, and I'm a strong believer in asking to the experts, gathering them, work together to clarify doubts and procedural issues to shorten waste reading.

Best regards and have a nice day.

Hiram.

Severini, M. F., Buzzi, N. S., López, A. F., Colombo, C. V., Sartor, G. C., Rimondino, G. N., & Truchet, D. M. (2020). Chemical composition and abundance of microplastics in the muscle of commercial shrimp Pleoticus muelleri at an impacted coastal environment (Southwestern Atlantic). Marine Pollution Bulletin, 161, 111700.

Valencia-Castañeda, G., Ibáñez-Aguirre, K., Rebolledo, U.A. et al. Microplastic contamination in wild shrimp Litopenaeus vannamei from the Huizache-Caimanero Coastal lagoon, SE Gulf of California. Bull Environ Contam Toxicol 109, 425–430 (2022). https://doi.org/10.1007/s00128-022-03568-6

Hello Hiram Herrera. Thank you for the clarification, it sounds like an interesting project and, based on what you described, muscle seems to be the most appropriate for the question that you are asking. If MPs are in muscle tissue then they are almost assuredly to be in the digestive system. If ingestion is the primary route of exposure then it would be interesting to compare particle size in gut and muscle to determine sizes that are able to pass the gut wall. But that may be outside of the scope of your project.

Visual sorting is certainly an acceptable and commonly used method for MPs in various types of media. Most people do visual sorting after some sort of separation (density separation or filtration), which it sounds like you have done already. The size MPs that you capture depends on things like the pore size of your filter. Visual examination is usually the next step. While not a firm threshold, ≥500 um is generally regarded as the size particles where manual sorting is the most effective. That doesn’t mean that you can’t sort out smaller ones if you can see them with your microscope, but the error rate increases as the particle size decreases. Visual sorting is prone to giving false positives because it relies on your judgement. And while a trained, experienced person has a lower error rate, this rate is still not zero. This is why a subsequent polymer analysis or staining is necessary. Nile red staining is easy, cheap, and fast especially if you have a lot of samples. Autofluorescence can occur in different amounts in different types of tissues, even within the same animal. Endogenous tissue elements or structures (e.g., collagen, bile) will fluoresce at certain wavelengths, it is a natural emission of light from structures when they absorb light. If the wavelength of this emitted light is within the same range as that of your stain then it can be more difficult to differentiate your stained object from these natural components. I don’t know if shrimp muscle tissue does this, but you will know when you put filters from procedural controls (or uncontaminated tissue) on the same microscope.

Classification (size, color, shape) is doable. For the very small MPs, it again relies on the resolution of your microscope. I don’t know of a standardized procedure that describes when you should stop looking for them in your sample. We usually just stop when we think we have found all of them. Unfortunately, there isn’t really a consensus on size categories (e.g., micro- vs. nano-) and even color. I have taken pictures of MPs and then used ImageJ (free software) to measure their sizes, and then either lumped data together into size categories/ranges or created a distribution curve. Color is often kept to basics: red, green, blue, black, white, transparent, multicolored, and other. But you could divide it up even more. Definitions for shapes (fragment, fiber, bead, pellet, film, etc.) are more agreed upon. I would recommend reading Hartmann et al (2019) for a good discussion on these classifications. Now, numbers/quantity is a different story. If you have a small number of MPs in your sample then you can count them all directly. If you have a lot of particles then you may want to employ some sort of grid method - you can google that and get lots of ideas - and then normalize to amount (wet weight, dry weight, volume, etc.) of starting tissue sample.

Since you are still in the methods development stage, you could also consider trying a protocol validation. Take clean shrimp tissue and spike it with different sizes, quantities, and/or types of MPs and see if you can get them back using the methods you have selected. It is a little time consuming and not necessary, but it could help.

- Melissa

Hartmann, N.B., et al., 2019. Are we speaking the same language? Recommendations for a definition and categorization framework for plastic debris. Environmental Science & Technology 53, 1039-1047. http://doi.org/10.1021/acs.est.8b05297

Melissa Chernick Thanks so much for your help. Your recommendations will be of great guidance. I don't have enough words that express my gratitude for all the information and time you provide and took to answer my questions, even when I know some of them were sort of basic. With all of this in mind, I think it wouldn't be of such harm to get into some kind of MPs identification/processing course and read a LOT more than I have so far :P. Now it's only matter of get organized and make things happen. Thank you again, Melissa, it was very very kind of you. Greetings and best wishes!

Hiram.

Microplastics are small plastic particles that are less than 5 mm in size, and they can be difficult to detect and measure due to their small size. The particle size limit for visual sorting of microplastics will depend on the specific method being used and the equipment and resources available.

Some methods for visual sorting of microplastics involve using microscopes or other magnifying equipment to view the particles, which can allow for the detection of particles down to a few micrometers in size. Other methods involve using sieves or other mechanical separation techniques to separate the microplastics from other materials, and these methods may have a larger particle size limit, typically in the range of 1-5 mm.

It is important to note that visual sorting is only one of many methods that can be used to detect and measure microplastics, and it may not be suitable for all types of samples or applications. Other methods, such as chemical analysis or imaging techniques, may be needed to accurately measure and quantify microplastics in certain situations.

It is also important to carefully consider the limitations and potential biases of each method when using it to study microplastics, and to use a combination of methods whenever possible to ensure the accuracy and reliability of the results.