Science topic

Crustacea - Science topic

Crustaceans form a very large group of arthropods, usually treated as a subphylum, which includes such familiar animals as crabs, lobsters, crayfish, shrimp, krill and barnacles. The scientific study of crustaceans is known as carcinology (alternatively, malacostracology, crustaceology or crustalogy).
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Is this speciemen probably a barnacle? It has been seen with orbitolinids of Early Cretaceous.
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Dear Prof. Ferre
What are the chances for this speciemen to be fragments of a blastoid. In Boardman, Cheetham Fossil Invertebrates, page 582, Figure 18.37, Hydrospire in a blastoid is depicted, which shows some resemblance to upper part of this speciemen, I would be thankful to have your response.
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Hellow , I need your help in detecting a universal primer for moleculer study of ostracod, who has information about it tell me please.
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Thanksfull dear
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Hi,
I am looking for papers on electrical stunning in crustaceans where the pros and cons of dry versus wet stunning is discussed / described.
Thanks.
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Electrical stunning is a method that is used to humanely kill crustaceans, such as shrimp and lobsters, prior to processing or cooking. There are two main types of electrical stunning: dry electrical stunning and wet electrical stunning. Both methods have pros and cons, and the most appropriate method to use will depend on the specific circumstances and the goals of the processor.
Dry electrical stunning involves passing an electric current through the crustacean's body while it is dry, typically using electrodes that are placed on the body or in water. This method has the advantage of being relatively simple and inexpensive, and it can be effective at rendering the crustacean unconscious. However, dry electrical stunning can be less reliable than wet electrical stunning, and there is a risk of overstunning or understunning the crustacean, which can result in animal suffering or reduced meat quality.
Wet electrical stunning involves immersing the crustacean in a conductive solution, such as seawater, and passing an electric current through the solution. This method has the advantage of being more reliable and consistent than dry electrical stunning, as the conductive solution helps to evenly distribute the electric current throughout the body of the crustacean. Wet electrical stunning is also less likely to cause damage to the meat, as it does not involve the use of electrodes. However, wet electrical stunning requires specialized equipment and is more expensive and labor-intensive than dry electrical stunning.
Both dry and wet electrical stunning methods have the potential to cause animal suffering if not properly implemented, and it is important to follow proper guidelines and protocols to ensure the humane treatment of crustaceans. It is also important to consider the economic and practical considerations when choosing between dry and wet electrical stunning methods.
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Hi everyone! I'm trying to identified this species, I think that belong to petruca panamensis but no so sure
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Thank you for your help John. I'm going to write to him and take more photos as well
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Hello,
I am a graduate student researching the diet of Green-winged Teal in North Carolina. We collected gizzards from 51 hunter-harvested teal this past winter, and I have just finished analyzing the contents. The unknown food item in question has appeared in 43 of 51 gizzards (84%), and most gizzards only had 1-10 of them but one had upwards of 80 in a single gizzard. I'm not sure what they are-photos attached. They are tiny, 1-2mm in length, and resemble tiny claws or mandibles with serrations. They seem chitinous or sclerotized, which is maybe why they have resisted being crushed up by the gizzards and are more prevalent than other food items. I emailed an entomologist at my university who said they aren't mandibles, but might be part of a plant, or some invertebrate with only a few sclerotized parts. The teal were in coastal North Carolina, so they roosted overnight in coastal saltmarsh dominated by cordgrass, and then spent the day in freshwater flooded cornfields. I have a feeling that they could be some kind of micro-crustacean they consume while in the saltmarsh. If anyone could eliminate possibilities or provide some insight it would be greatly appreciated. Thank you!
-Cole Tiemann
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Have the https://ucedna.com/ cal-e-dna run the DNA test, for less than $200
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i got this crustacean can anyone tell if its benthic isopod?
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Gnathiidae is probably correct -- can a brief account of its habitat be provided?
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I'm asking this question to know if it is possible to rely on carapace shade/color in order to determine the animal's taxonomical group.
Thank you in advance for your answer.
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Not my subject, but I think the shadow can be a distinction between species in a population.
In other words, an aid to sorting the individuals in a population. Absolute identification may require other criteria. Unfortunately, keys are often difficult to use. Once the species is known, one can often recognize it on the habitus.
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I work on ostracod species that belong to order crustacia and phylum Arthropoda but I dont have enough key for this ,please who can help me
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Hello ! If you're working on inland aquatic Ostracods I invite you to check this document :
P.S.: Please bear in mind that it isn't for species level identification but it's a really good starting point.
Have a great day !
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I am running this code (below) to add error bars onto my bar chart, i am trying to to work out the standard deviation (sd) of shannons diversity index
The code works up until the second/third last line
geom_errorbar(aes(ymin=shannon-sd, ymax=shannon+sd), width=0.2,
position=position_dodge(0.9))
This error code keeps coming up
"Error in shannon - sd : non-numeric argument to binary operator"
but this is my data...
Exposure genus shannon sd
Exposed Crustacean 0.000000 0.00000000
Exposed Mollusc 1.199625 0.13291129
Exposed Seaweed 1.513125 0.42093822
Sheltered Crustacean 0.025500 0.07212489
Sheltered Mollusc 1.156750 0.26763341
Sheltered Seaweed 1.848125 0.27264128
anyone know where im going wrong?
library(ggplot2)
#+++++++++++++++++++++++++
# Function to calculate the mean and the standard deviation
# for each group
#+++++++++++++++++++++++++
# data : a data frame
# varname : the name of a column containing the variable
#to be summariezed
# groupnames : vector of column names to be used as
# grouping variables
data_summary <- function(data, varname, groupnames){
require(plyr)
summary_func <- function(x, col){
c(mean = mean(x[[col]], na.rm=TRUE),
sd = sd(x[[col]], na.rm=TRUE))
}
data_sum<-ddply(data, groupnames, .fun=summary_func,
varname)
data_sum <- rename(data_sum, c("mean" = varname))
return(data_sum)
}
df3 <- data_summary(diversity, varname="shannon",
groupnames=c("Exposure", "genus"))
# Convert dose to a factor variable
df3$genus=as.factor(df3$genus)
head(df3)
p <- ggplot(diversity, aes(x=genus, y=shannon, fill = location)) +
geom_bar(stat="identity", width = 0.5, position=position_dodge()) + theme_minimal() +
geom_errorbar(aes(ymin=shannon-sd, ymax=shannon+sd), width=0.2,
position=position_dodge(0.9))
p + scale_fill_brewer(palette="Paired") + theme_minimal()
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Could you read the data into a very simple program, possibly multiplying the numbers to see if you get the error because the file is corrupted a bit. You could also move the lines around before running the program to see where it has the problem. As Liang Chen suggests, it seems like a simple problem with the file.
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I'm doing a thesis that is related to sound production of crabs. Various crabs such as Ghost crabs and Dungeness crabs produce sound when they feel vulnerable. In my case, my thesis focuses on the mangrove crab Scylla Serrata. But due to travel restrictions due to Covid-19 situation I'm not able to perform my own experiments. I could not find any resources that were dedicated to sound production of Scylla Serrata. So, I wanted to know if Scylla Serrata produce sound when they feel vulnerable or before other crabs cannibalize them.
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In the literature I found the following two interpretations:
- it is the maxillipedal exopod
- it is a specialized part of the maxillipedal epipod
So far, however, I found no real discussion giving arguments for one or the other. Is anyone aware of something like this?
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Maxillipeds are appendages that have been adapted to serve as mouthparts. These can be quite similar to pereiopods, especially in less mature decapods. Pereiopods have walking legs that are also employed for food collection.
Maxillipeds. The first three pairs of legs are transformed to maxillipeds, which help in food manipulation by passing food forward to the mandibles for chewing or to the maxillae for cutting into smaller pieces.
Maxillipeds are appendages that have been adapted to serve as mouthparts. These can be quite similar to pereiopods, especially in less mature decapods. Pereiopods have walking legs that are also employed for food collection.
Maxilla is one of the two bones that make up the upper jaw, whereas maxilliped is one of the feeding appendages on the heads of certain crustaceans.
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I believe the class is crustacea but any help would be appreciated! This species was collected in Belize. Approximate size is 5mm
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Yes, it is an amphipod. My guess would be family Colomastigidae. Best is to contact Sarah Lecroy, she might know the critter:
All the best, Oliver
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> Animals are kept in accordance with ABNT standards.
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Algae
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I would like to know the age of some Excirolana armata individuals sampled. Is there any bodily structure or anything like that which is indicative of age in this species or family?
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Hello. All soecies of crustaceans growth after of ecdycis and no there arent structures with rings of growth, however, is possible with the measurement of length body use some method as battacharya.
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I already published some works on predation by drills on ostracods, however I wonder if there are recent publications on the subject!
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Thanks so much to you too, Prof. Otar Shainidze.
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For part of my research I am attempting to assess the abundance and diversity of crustaceans in an aquatic habitat. I intend to take picture of the specimens once collected before they are preserved and lose their colour. I mainly wanted to know if there were any specific guideline to taking taxonomic photographs of shrimp e.g. how it should positioned/oriented, should the appendages be positioned in a specific way as well?
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Hi Maizah, I concur with James: each species has a different set of characters, so you will have to study them in advance. I prefer to start taking photo's of living specimens, because they show some behaviour I want to register (see https://nieuwewendingproducties.blogspot.com/2018/03/in-vitro-in-natura.html - http://micksmarinebiology.blogspot.com/2017/10/spookkreeften-determinatietabel.html)
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Dear colleagues!
Have you ever seen a program which is able to select the most suitable pairs of group-specific primers from a set (more than 5 pairs) of proposed ones?
Suppose in my lab I have several pairs of primers for different groups of organisms (polychaetes, fish, mollusks, echinoderms) and I want, instead of ordering new ones, to select from them those that are absolutely accidentally found to be specific to... crustaceans.
In other words, a bulk search among available primers that have never been tested for specificity to a particular group.
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In general, everything can be found on
I did this for different types of beetles. The point is that you need patience, because if there are many records in the database, they must be sorted. On the site you can find sections of genes that have not yet been identified.
Regards, Sergey
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Chordotonal organs are found in insects and crustaceans but do they also occur in mites. Specifically in Phytoseiidae.
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Our knowledge of mites is poor in that aspect. There are soft tissue areas on the surface of the mites that we do not know if they do have such function, areas that look like possible ocular function, or perhaps vibration sensor. The correct answer is we do not know, but we should look at it as we were able to capture drop and stop behaviors of plant-feeding mites before the predator mite was over them. Many of these strange areas over the surface have been noticed by the use of the cryo-SEM. See also Functional morphology of tarsal adhesive pads and attachment ability in ticks Ixodes ricinus(Arachnida, Acari, Ixodidae)Dagmar Voigt*,‡and Stanislav Gorb 2017.
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My question is in relation to new trace fossils from Pleistocene-Holocene marino-marginal depositional environments with records of molluskan and crustacean burrows. In these outcrops the friable sediment, already partially cracked, makes impracticable the collection of most specimens, if any. However, the morphological details are faithful when compared with modern analogous or the same producer of these traces and their ichnotaxonomic importance is high. What are possible solutions to this issue?
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Trace fossils followed the same approach as the Code of Zoologic Nomenclature for body fossils, using the Linnaean binomial nomenclature (genus species). To establish a new genus or species requires a type specimen that represents the new trace fossil. This system is imperfect at best and not all that useful, but it has the momentum of history. Photos would not be an accepted basis for naming a new trace fossil. If it is difficult to collect you have no objective evidence for others to compare it with, but if you know what animal made the trace you are farther along than most ichnologists will ever be.
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Two weeks ago I found a peculiar crab in the intertidal zone in SW Florida. when found, the crab was not utilizing a shell. It was similar shaped to porcelain crabs found in our area, but with smaller claws (red tipped) and the 5th perepod was flipped up on the dorsal size of the carapace. The eyes were also uniquely positioned on the ventral side of the animal. Upon collection, the crab was observed carrying a cockle shell half. It fit particularly snug in this shell, which leads me to believe this is a commonly used shell for this species. Any assistance on identification and/or resources would be appreciated.
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This is, without a doubt, a Shellback crab. Genius : Hypoconcha.
It seems like a Hypoconcha arcuata.
There is two more species in Florida (Hypoconcha parasitica and Hypoconcha spinosinima).
Whichever species it is, it's a beautiful speciment!
Cheers,
Greg
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these crustacean found in a gut content of Ostorinchus margaritophorus (Apogonidae)
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Halichoeres argus is a marine wrasse; reef-associated (https://www.fishbase.se/summary/Halichoeres-argus.html);. By the way, wrasses, which are found in the Indo-Pacific, frequent all coral reefs, so to speak. It is among them that Wrasses which perform cleaning services are encountred. Your observations lead to ask the following question: is the wrasse Halichoeres argus a cleaner fish? To answer this question, an exhaustive bibliographical research is necessary as well as a study of the stomach contents of this species statistically valid in time and in space.
Moreover, most wrasses are carnivorous predators and eat small crustaceans, snails and worms which would explain the presence of other free Crustaceans as Peltidium sp. (Crustacea, Copepoda, Harpacticoida) in the gut content of Halichoeres argus which would be a complement to the cleaning activity.
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I'm PhD graduated in fisheries. Have some experiences in crustaceans propagation, trout egg hatchery, larval rearing in RAS system, aquaculture in the floating HDPE cages in Caspian Sea and etc.
You can find my curriculum vitae in the attachment. All my fisheries research and activities are mentioned there, in brief.
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I am looking for a funded PhD position in Aquaculture/Fisheries. Preferred locations are Germany and Canada. I have an MSc. Fisheries Science from Universiti Putra Malaysia and a B. Fisheries and Aquaculture from University of Agriculture Makurdi Nigeria.
My research interests are in the areas of RAS, Aquaponics, Biofloc and wastewater aquaculture.
Kindly find my CV attached for your reference.
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We found this small, maybe parasitic, crustacean in the North Sea offshore. The posterior body part is missing, but it looks quite characteristic. There is double-hooked rostrum, 2 antennae (second pair broken off), 2 small eyes on each side of head, no coxal plates, and maybe a sucker-like structure near mouthparts.
Stained with Shirlastain A.
We don't have any clue to which higher group it belongs, and would be happy if anyone can give a hint.
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This is the calanoid copepod Anomalocera patersoni, which is a regular inhabitant of the North Sea plankton:
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I am trying to ID fairy shrimp but am unable to find any lists or keys to use.
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Jiri, you might have luck with this outdated guide:
Belk, D. 1975. Key to the Anostraca (fairy shrimps) of North America. The
Southwestern Naturalist 20 (1): 91-103
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Can anyone identify this species? It's from a temperate lagoon located in the southern part of the Po River delta (Italy).
Many thanks,
Monia
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It is an Anthuroidea species for sure. The estuarine character of your sampling location indicated the possibiloty of Cyathura carinata, but it can also be another species.
Please check Wägele (1981) Study of the Anthuridae (Crustacea: Isopoda: Anthuridae) from the Mediterranean and the Red Sea. Israel Journal of Zoology 30: 113-159
Cheers
Michael
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In looking for a different treatment for small crustaceans that parasites sharks and rays
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I don't have any experience with Cyromazine but diflubenzuron (trade name Dimilin) is commonly used in public aquaria, as is trichlorfon (Trade name Dylox) - though dylox is an organophosphate and really nasty stuff to work with, also rays and skates tend not to tolerate it as well as dimilin. Milbemycin oxime, commonly used with dogs and cats under the trade name Interceptor will also target crustaceans, and is so benign it is even commonly used in aquaria with corals and jellyfishes.
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I need to differentiate living veligers from dead. Is there some staining or another method?
Thank you for your help in advance
MAria
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Javier Morales la idea era poder distinguir tras un tratamiento para la eliminación de larvas, si era o no era eficaz. Por ello tras el tratamiento se tomaban muestra y se evaluaba el % de larvas muertas y de larvas vivas.
Los moluscos muertos si estaban fragmentados si que se teñian, pero no si no estaban dañados.
Un saludo
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Hi all, I will appreciate if you could direct me to a paper where I can find the stomach pH in crustaceans. I am sure it will vary between species, yet any data will be useful and much appreciated.
Thanks
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It is depend on what time of animal digestibility or other living activity but around neutral pH.
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Hello, I am a PhD student interested in population genetics of marine invertebrates. Currently I'm working with a bryozoan: Reteporella.
I have been trying to amplify COI (at least) and, although I finally got some bands, after Sanger sequencing, they turned out to be of bacteria, protists, or crustacea (parasits). I'm working with recent samples (collected between 2018-2019), and also some from 2010 (even got bands in these ones, but no sequence). I have been following the conditions suggested in papers dealing with Reteporella and other bryoans, using Platinum Taq (Thermo Fisher) ou Multiplex PCR Master Mix (Qiagen):
  • Folmer's LCO1490/HCO2198 universal primers
  • 94ºC 3 min; 40x(94ºC 30s, 45ºC 30s, 72ºC 1min); 72ºC 10min
  • 95ºC 5 min, 35x(95ºC 40s, 45ºC 45s, 72ºC 1min); 72ºC 8min
I'm aware these temperatures are quite low and prone to amplify inespecific targets. I'll try a gradient PCR 45-55ºC and a touchdown between the same temperatures.Can someone help me, any tips to help me get specific sequences of Reteporella?
Samples were extracted with PureLink extraction kit (Invitrogen), reccomended for "difficult-samples) after completely crushing the sample with a stainless steel pestle (which is passed on ethanol and fire between samples). Only samples that looked "clean" on the surface with polyps at sight (ensuring they were alive when collected) were extracted.
Thanks in advance!
*Attached are eletrophoreses of the PCRs that worked with annealing temperature of 45ºC using the MM Qiagen or Platinum.
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Thanks everyone for the very useful tips! I will have them into consideration when I encouter problems again.
I believe the main problem to be the universitality of the primers that were making everything difficult.
Paul Rutland Increasing the Ta inhibited the amplification, I got no bands at all.
Sibnarayan Datta The primers' stocks and diluted aliquots were new and working perfectly with gastropod taxa.
Heather E Doherty the long gel run is a great tip, thanks, I will dot it if the problem persists!
Angel del Marco i did not know SnapGene, I have been using Primer Blast tool to do a first evaluation of the primers vs taxa in study. I am using a commercial kit for total DNA as most of the samples are completely destroyed in the process and we are also using them to obtain nuclear sequences and SSR.
I eventually ordered COI primers specific for bryozoans, designed and published by colleagues, and the gels now seem more promising, I'm waiting for the sequencing results now.
Thanks everyone!
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Micro-Computed tomography (micro-CT or μCT) is a microstructural, non-destructive study technique that allows you to study the internal and external anatomy of biological samples, and perform their reconstruction through a virtual three-dimensional model.
he possibility for combining μCT with other techniques is one of the major advantages of μCT scanning, and the technical development of higher resolutions in lab-based μCT-scanners allows for investigating the anatomy of specimens in the sub-milimeter range See:
Krieger, J., & Spitzner, F. (2020). X-Ray Microscopy of the Larval Crustacean Brain. In Brain Development (pp. 253-270). Humana, New York, NY.
and:
KHAN, M., WHITTINGTON, C., THOMPSON, M., & BYRNE, M. (2019). Arrangement and size variation of intra-gonadal offspring in a viviparous asterinid sea star. Zoosymposia, 15(1), 71-82.
This technique would allow us to explore how different structures develop in species that develop their life cycles in different areas with different characteristics and find a relationship between them.
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Except that the radiation dose may be excessive in the study of the biological object (presumably alive) that you are studying especially at higher resolutions. And, that there is potentially poor differentiation of soft tissues using CT. Other small animal imaging modalities, ultrasound, MRI, etc. may offer similar useful and complementary information.
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For a long time we are looking for the fossil cladocerans (Crustacea: Branchiopoda: Cladocera). Many papers were published previously by my team. But now we are looking for the next fossils, i.e. in Mesozoic rocks, any amber pieces etc. Did you see them?
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I think you mean 'interested' lol
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why moulting process in crustacea
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Hello Khaled; The exoskeleton of arthropods is inelastic - the animals' container is rigid. Thus the animals can't grow unless they periodically produce a new, larger exoskeleton. Best regards, Jim Des Lauriers
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Im currently working on gut content analysis of some juveniles from seagrass bed in Seribu Archipelago, Indonesia. I found some difficulties as most diets were found in not complete part and perhaps broken. Thank you
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First of all in your in your question you failed to specify whose juveniles you are working with. Still,based on the five photos, without considering the others attached -
The calanoid copepods are either partially digested and the cyclopoid (may be Eucyclops)copepod is in its copepodite stage and also there exists a nematode worm.
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In transcriptomic analysis of crayfish infected by a pathogen ( oomycete), we found in hemolymph, chitinases and Lpmos... is it a general defence mechanism? thank for your help
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If we consider the "general defence mechanism" of crustaceans, it primarily involves beta-glucan-binding protein, lipopolysaccharide- and glucan-binding protein (LGBP) and lectins produced by various haemocytes. These pattern recognition proteins activate prophenoloxidase (proPO) system to initiate immune response. LGBP is the main molecule in response to infection with fungi and oomycetes.
As to your observation, it can be a tricky business because it is not clear whether the chitinases and LPMOs are from the crayfish or oomycete. This is because these chitin-binding molecules are actually the primary products of the pathogens. And particularly, LPMOs can be the pathogenic factors against the haemocytes of crayfish.
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I'm using Chitosan flakes, obtained from crustaceans. In my application, I put the chitosan flakes with caustic soda so that the pH is over 13. Now my question... Does the high pH not lead to additional deacetylation? However, IR measurements on different deacetylated chitosans have shown that there are no significant differences via pH treatment. The high pH does not lead to further deacetylation. Why is that? Why is virtually no 100% deacetylation possible?
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Bitte mal den Kollegen kontaktieren und ihm diese Frage stellen
MfG
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Dear Colleagues, we would be most grateful if you could complete this very short survey on Endocrine Disrupting Chemicals (EDCs). This survey is intended to inform the wider scientific community about the progress and impediments to endocrine disrupter research in invertebrates. We welcome views from those working outside invertebrate toxicology
Please follow this link https://forms.gle/tZAbHnhn6fArnAZR8
By participating you will be giving consent to your anonymous data entries being used as part of the survey. Due to the anonymous nature of this survey participation can't be removed. For further details on this project and how the data will be used please feel free to contact alex.ford(@)port.ac.uk [Please remove the () around the @]
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Hello ;
Dear Alex
Several effects of PE on invertebrates
Disturbances of larval stages
Egg malformation
Disruption of the nervous system at the target level; receptors
Modification of biosynthesis; metabolism and elimination of natral hormones
Risk for the species and the higher-level food chain by bioacumulation of persistent substances
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I would like to know which DNA extraction kit would work best for extracting bacterial DNA from animal tissue, specifically the pleopods of a crustacean. Some people have mentioned the DNeasy blood and tissue kit has worked for them, while others have suggested the power soil kit. I understand a kit with beads to aid in bacterial cell lysis is preferable, but any advice would be appreciated.
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With all respect to my colleagues Reza Hadavi and Alireza Mordadi The DNeasy Quiagen Kit is not the best for this application. The DNeasy kit relies on PBS and proteinase K to lyse cells and this is not efficient for bacteria nor is it sufficiently harsh to recover DNA from bacterial cells that are adherent to the pleopod surface. The Kit I mentioned above uses a strong denaturing solution to lyse the sample and will be superior for extraction of DNA from bacterial cells and hard tissues such as crustacean exoskeleton. In fact, the paper cited by Reza Hadavi shows that a genomic DNA isolation kit similar to the one I mentioned above performed as well as the Qiagen kit. While the Qiagen kit is an excellent product and I have used it myself, as someone who has experience in extracting DNA from arthropod samples specifically I can tell you that it will not be as effective in this application. The paper cited is extracting DNA from bovine soft tissue, and that is a much different matrix than crustacean pleopods. April Stabbins original question was what method is best for extraction of DNA from crustacean pleopod tissue. Simply stating that a variety of techniques exist or recommending a kit or method that hasn't been validated for the application in question is not helpful. Please kindly answer questions about which you have genuine expertise and in a manner that addresses the issue of the original poster.
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Hi. I would like to ask you.
We typically fix fish tissue with 4% PFA and 30% sugar for FISH or IHC.
Recently, however, the failure of frozen section for FISH or IHC using shrimp, a crustacean, has been continuing.
Does shrimp have to be fixed in other ways?
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Typically seawater buffered Davidson's or 95% ethanol. Shields (2017) provides extensive discussion on differential fixation: Jeffrey D Shields, Collection techniques for the analyses of pathogens in crustaceans, Journal of Crustacean Biology, Volume 37, Issue 6, November 2017, Pages 753–763, https://doi.org/10.1093/jcbiol/rux077
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Instantaneously?! For growth, the new exoskeleton is soft and pleated under the old exoskeleton. When it molts, the animal inflates (with water or air), gets out of its old shell and unfolds its new exoskeleton to grow. But this shell is incomplete and have to be completed by complementary secretions. More, the natural sclerotisation time of the new exoskeleton is not instantaneous because it involves various chemical processes: Calcification using external product (crustacean), or tanning (insect).
Alternatively, for instantaneousness in biology, you could contact Harry Potter ;-)
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Currently working on marine macro-invertebrates (especially molluscs, echinoderms and crustaceans) from gleaners' catch for my thesis and need help in taxonomic identification and confirmation. Thank you.
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@ Jyl C Marfil : You are welcome
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Dear all, 
We’re currently extracting total RNA from gammaridean amphipods (Gammarus pulex). Animals were freshly collected, directly frozen at -80°C.
Trying different RNA extraction protocols (Qiagen, Trizol+Qiagen) we always end up seeing no obvious 28S band. First we thought it is due to the denaturing step prior to loading onto the Fragment Analyzer (known issue). But omitting this heat denaturing step does not lead to a change - the picture remains (see attachment for examples). Any ideas? We also gave the samples to a different lab - extraction results the same. There seem to be sometimes two only marginally different peaks at the 18S band position and we suspect that something other than heat is leading to the dissociation of the two 28S subunits possible. If these are of same lengths as the shorter 18S fragment can this explain the pattern… So non-model organisms experts: Can someone help us with this puzzle? Best Florian and Team
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Hi!
I am preparing ostracods for identification and I need to open the valves. Is there a best way of doing it?
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Thank you all for the very helpful responses!!
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My scientific advisor (Burukovsky R.N.) is a famous shrimp researcher and carcinologist. He analyzes food from stomachs of many crustaceans. So... it is an obvious situation when he found an interesting unidentified object inside. Somebody can help us and tell something about what is it? Small bonus below: the book about shrimp feeding in Russian. See foto too. Size of object 0,5-1,5 mm. My E-mail: echo_tc2@rambler.ru
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An algal specimen possibly from Dinoplagellates. Planktonic algae normally act as preferred food of Crustacea. The specimen however, needs thorough examination under microscope to achieve taxonomic precision in its identification.
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I'm trying to extract the RNA from the shrimp optical ganglion.
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I don't know.
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Crustaceans blood colour.
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In Invertebrates, the hemolymph, that is to say the liquid that plays the role of blood, does not contain hemoglobin (as in Vertebrates) but hemocyanin. This oxygen carrier pigment contains copper atoms instead of iron atoms (as in hemoglobin), which gives it a blue-green color.
It is for this reason that the blood of insects or crustaceans is not red, but blue-green. The best-known example of a blue-blooded animal is that of an arthropod sometimes called "blue-blood crab": the horseshoe crab (Limulus sp.).
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Hey everyone,
I was wondering if anyone has tested/published some crustacean focussed 18S primers that provide an amplicon size of around 1kb to use for phylogenetics?
Many thanks!
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Hi,
I am working on the screening of Crustacean Viruses. Meanwhile, when I screened my samples for IHHNV (amplicon size 309bp) my positive control is not working. I am getting multiple bands just below the well (gel image attached, last lane is the positive control).
Can anyone please suggest, why is it happening so?
I have carefully done conducted the experiment and repeated too, but still the problem is the same.
Please help me out in solving these issues.
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What is the species that is moving around the echinoderms larvae?
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Hi Francisco
It is normal to find ciliates or another protozoans organisms living with some larval stages of larger invertebrates. These tiny organisms could be flagellates.
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At low salinity (below 4 ppt), the proportion of major minerals (Cl, Na, SO4, Mg, Ca, K) is important as well as the absolute concentration for the growth and survival of marine shrimps. Does anyone know this statement still works in brackish water (10-25 ppt)? And what mineral is the most important?
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Dear all,
I have gathered samples of Dreissenid mussels at different locations in a lake. After counting and measuring the mussels, histograms featuring the number of individuals per shell length ranging from ≤0.4cm to 3.5cm have been created, most of which are showing multimodal distributions but some are unimodal, too.
What is being tried to do, apart from visually determining the size-frequency distributions, is to apply a statistical mathematical tool in R that goes through all of these data and attempts to classify recurring groups that then ideally represent age-groups (cohorts, populations) of the mussels.
It seems like packages in R that might be helpful are 'mclust' and 'Rmixmod' both of which have already been tried. However, in the end, always a dead-end has been reached so it got me wondering whether the arrangement of my data may be the problem or there is another underlying cause.
Has anyone already encountered and maybe solved equal problems and might possibly be up to having a look into the organisation of my size-frequency data?
Similar methods to what I am trying to do have been exercised for example in:
Comtet, Desbruyeres (1998) Population structure and recruitment in mytilid bivalves from the Lucky Strike and Menez Gwen hydrothermal vent fields (37'17'N and 37"501N on the Mid-Atlantic Ridge)
and
Taylor et al (2009) Using length-frequency data to elucidate the population dynamics of Argulus foliaceus (Crustacea: Branchiura)
Thanks to everyone wanting to help!
Benjamin Wegner
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Hi, you can have a look at the R library "TropFishR" in R-cran (https://cran.r-project.org/web/packages/TropFishR) or in github (https://github.com/tokami/TropFishR). The Bhattacharya’s method can suit your problem.
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I want to extract chitosan from crustaceans such as shrimps and lobsters. How would this be possible in the simplest way?
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Hi Cemal,
To synthesis chitosan from crustaceans, follow these steps
1. Cleaning: Remove the shells from the shrimp or lobster, clean well, dry and blend it.
2. Decalcification: using aqueous HCl (3 to 5% w/v) at room temperature overnight.
3. Deproteinization: using aqueous dil. NaOH(3 to 5% w/v) at room temperature overnight or 80 to 200 oC for few hours. You will get the product CHITIN
4. Decolorization: using 0.5 % KMnO4 and Oxalic acid. (You can avoid this step if you use colourless endoskeleton such as squid gladius)
5. Deacetylation: using hot con. NaOH (40 to 50% w/v) at room temperature. Now you will get an end product CHITOSAN.
Check this links for more information:
Thank you.
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The aims is for detect the hormone profiles in crustacean's hemolymph. Therefore, before proceed with the GC-MS procedures, the hemolymph need to be dissolve with derivatives. Does anyone can help me to identify the suitable derivatives? Based on my experience with ELISA, methanol and ethyl ether can become the derivates. But I am not sure the suitable derivatives for detecting hormone profiles by using GC-MS procedures. Thank you.
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Check this link of article.
Greeting
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is anybody familiar with species identification of Euphausiidae shrimps, also known as krill? A co-worker (W. Langbroek) collected a specimen in a shallow marine bay along our Dutch coast at Katwijk (52°12'39.74"N , 4°24'0.41"E on 24.iii.2018) and I only have the key by Mauchline (1984). In The Netherland, 2 species have ever been reported i.e. Meganyctiphanes norvegica and Nyctiphanes couchi. My specimen is a bit small (about 9 mm) so perhaps not all features characteristic for those species are fully developed. As lateral denticles are missing it seems no M. norvegica, but also a comb-like structure on antenna 2 is missing characteristic for N.couchi. However, I think that it may still be a juvenile specimen of the latter for it has 7 pairs of legs with the last one very much reduced, eyes are undivided, no lateral denticles on the carapax and with reflected lappet on the first antennal segment. Do you agree? Or do you have another suggestion? Are there any other keys for Euphausiid crustaceans which can be used for the NE-Atlantic?
many thanks in advance
Ton
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note the middorsal spine in N. couchi
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I have seen ectocommensal triclads and their egg cases in Antarctic isopods (undescribed to the best of my knowledge), Bdelloura species in horseshoe crabs.. I wonder if there are potentially triclads in other crustacean species?
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Thank you!!
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Hi Everybody,
I need ideas on methods which could be used to mark small shrimp (approximately 3 cm total length) for the purposes of identification of individuals in a lab-based study. It doesn't need to be easily discernible on a camera, but it does need to be easily discernible if the shrimp were to be captured out of its tank with a net and examined. I was going to use nail polish, but now I've read that it takes a very long time to dry. The less invasive the method of marking, the better, and the simpler and cheaper the method of marking, the better. Thanks in advance.
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As long as your shrimps are not moulting you can use superglue mixed with a some inert substance to give it colour and placed as a dot on their backs. I used this method to do a mark recapture experiment on Crangon crangon. Superglue dots will harden underwater.
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In fact, I'm looking for a way to maintain a good survival an growth of these shrimps, especially at early life stages (I've heard about some ion deficiencies in marine sea salt which lead to a higher mortality of larvae or juveniles).
Have you any advices concerning the salt quality?
Thanks in advance!
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If you are considering artificial salts, you are also using recycled aquaculture systems (RAS) so my response includes that assumption.
For maturation, spawning and larval rearing high-quality synthetic salt mixes are required that include all trace element and very high-performance RAS. Any water change to prevent the buildup of organics and Nitrate can be added to the growout stage waters.
After PL 10 or so, they become more water quality tolerant and that allows cheaper synthetics that just pay attention to Na/K, Ca/Mg, borates, and TDS, alkalinity etc. while depending upon feed for the input of a lot of the trace elements like Cu, Zn, Fe, Mn, F etc. Denitrification is required to minimize the salt cost.
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This relationship is very obvious to me (think of lobsters), but hard to actually measure. I could not find any study which addresses this issue!
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Not sure how to measure cuticle thickness per se, but you could use an indirect measure: how hard you have to press to puncture the carapace. John Christy and collaborators have done a similar thing a few years back for fiddler crabs (reference below), and they showed that it is harder to puncture the claw than it is to puncture the carapace. Also, the force you needed to puncture the claw was correlated to the pinching force of the claw. These suggest that Nahuel's idea is on the right track.
I would go a bit further and say that not only diet, but how much a given species fight should also correlate to cuticle thickness.
At the individual level I would assume that there should be a good amoung of variation as well. The minerals that make up the cuticle are not easy to come by in water, thus I would guess that some individuals can capture more minerals than others.
It would also be interesting to test if crabs that consistently win fights also have thicker cuticles - that may decrease the force the opponent is able to apply on the individual. That sort of study is missing in the fighting literature...
I hope some of my rambling is helpful.
Cheers,
Alexandre
The paper I commented is this:
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In decapod crustaceans, spermatophores studied to date have been categorized into three different types. One type of spermatophore is pedunculate and is present in all anomurans except for a few species in Hippidae (Greenwood, 1972; Trelli et al., 2007; Tudge, 1999; Scelzo et al., 2004). Another type of spermatophore is tubular and has several layers made of acellular material and they are found in the form of an interrupted column as in A. leptodactylus, Pacifastacus leniusculus, Homarus americanus, Enoplometopus occidentalis (J. W. Randall, 1840) and Panulirus homarus (Kooda-Cisco and Talbot, 1982; Haley, 1984; Mann, 1984; Radha and Subramoniam, 1985) or units pinched off the column (Dudenhausen and Talbot, 1983). The third type of spermatophore is the simplest form of spermatophore and is found in brachyuran crabs. They are small and either ellipsoid or spherical. This type might form the sperm plug in the seminal receptacle of the female that prevents other male sperm from getting into the female reproductive tract (Cronin, 1947; Ryan, 1967; Hinsch and Walker, 1974; Krol et al., 1992).
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Thakns dear Mariel
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I'm finishing an article of my research, which consists in managing some crustacean species names, but some of them went through taxonomic changes, specially genus level changes. One example is Palaemonetes pugio that was changed for Palaemon pugio, meaning a genus change and therefore a comb. nov. (or combination nova). But my problem relies on how to cite the authority of the species. the former name and authority goes like this: Palaemonetes pugio Holthuis, 1949... but now is Palaemon pugio and the reappraisal was made by De Grave and Ashelby (2013). In my paper I wrote: Palaemon pugio De Grave and Ashelby (2013) but one of my reviewers said that this is incorrect and wrote Palaemon pugio Holthuis, 1949, which means to maintain the current accepted name but the author of the original and former name. So I don't know exactly what to do... if anybody has had the same issue please help.
Thank you very much.
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Hiram:
As per Article 51.3 of ICZN, you may cite your new combination as follows:
Palaeomon pugio (Holthius, 1949) De Grave & Ashley, 2013.
Best
Syed
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I'm searching for some resource (book or paper or database) with more or less complete list of common marine zooplankton species (e.g. crustaceans) with body lengths or better body masses. There are many publications in this direction, but rarely raw data is presented. I would appreciate any advise.
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Dear Danilo,
Thanks a lot. It's closely related topic. I also found some publications on regressions. If it's of interest, there are some more recent examples:
Nakamura et al. 2017 "Length-weight Relationships and Chemical Composition of the Dominant Mesozooplankton Taxa/species in the Subarctic Pacific, with Special Reference to the Effect of Lipid Accumulation in Copepoda"
Robinson et al. 2010 "Length–weight relationships of 216 North Sea benthic invertebrates and fish"
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Permian system, Sakmarian stage. Russia, the Urals. Carapace
Phyllocaridan? Another Malacostraca?
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That looks very robust for a phyllocarid, to me. Could it be part of one of the early decapods?
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Dose anybody have some photos from below crab species? I could not find. 
Potamon gedrosianum
Potamon magnum
Potamon mesopotamicum
Potamon ruttneri 
Potamon strouhali
Potamon transcaspicum 
 Potamon bilobatum 
Potamon ibericum 
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You can have a photo of Potamon gedrosianum in this paper of mine
Records of non marine brachyuran crab species of Pakistan including note on their ecology & description of juveniles of Sartoriana blanfordi
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- Best quality and reliable ELISA kits.
- Journal / Article references used for crustacean allergen quantification.
- If possible approximate price/ cost of that ELISA kit.
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Dear Md Faisal,
I thought you might like to know that we just added another supplier to our Directory that also offers a Crustacean Tropomyosin ELISA. Here is their page:
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Recent discover of trilobite eggs has me wondering whether all or some trilobite species were single-birthers like some arachnids, crustaceans, and other plants and animals, albeit very limited.  Wondering whether phylogenetic analysis would be a starting point to support or refute the question.
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I think this is highly unlikely. The main reason is that most trilobite species moulted several times after reaching what is regarded as maturity: their final morphology, with adult number of thoracic ribs. There are also quite a few cases of suspected nesting behaviour, where pairs of the same species (at similar growth stage) are encountered within a shelter such as a nautiloid shell; I've seen this is Wales, and it is common at some levels in Bohemia.
The individuals involved in this are normally not the largest examples known, or even close to it. Therefore, if these were reproductive pairs, then they could survive to moult several more times. Unless reproduction was unlikely at an early adult stage, semelparous trilobites should never have got to their full size.  
This might not apply to all trilobites, of course, and my wife's just reminded me about agnostids... which seem like a good candidate for semelparity, if there is one!
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Hello, 
I have been passed this photo of a small crustacean (Malacostraca?) eating planted mangrove seedlings (Rhizophora) in central coastal Vietnam. They are girdling the seedlings, which then fail. 
My knowledge does seem to be limited to: oh, looks like a sea louse (!), so I am asking the question.
The situation is somewhat brackish evidently, due to fluvial influence at that part of the estuary, and there is no herbivory in more saline  water plantings.
The picture isn't great, but hopefully enough. I do not expect there are easy solutions to the pest problem in mass reforestation efforts!
(Planted seedlings do seem to be generally more susceptible to herbivory and I do think, without visiting the site, that the mangrove species selection may be wrong here, but that is another topic!).
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Yes, there is debate about nurseries for the Rhizophoraceae, especially Rhizophora because the 'propagule' (viviparous seedling) is so easy to direct plant. The VN experience in reforesting after the US defoliation in Rung Sat (Can Gio now) and Ca Mau and continuing now in farmers planting in aquaculture areas was/is direct sowing. So easy.
Generally nursery culture is a waste BUT perhaps not in this case if the advanced seedlings are more resistant if grown on in a nursery. An avenue to try, although substitution of the first planted species in affected areas to Avicennia or Sonneratia might be better!? 
Gazi Bay is more saline I think, both being pretty open to the ocean and not having the same intensity of wet season, where everything turns fresh as here, even out to sea at the mouths of the rivers! 
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Can you help to identify this crustacea?
Obtained from the Persian Gulf
Almost less than 20 mm size