Culture Journal, Species: Parvocalanus crassirostris

General
Species:  Parvocalanus crassirostris
Species description:  Calanoid copepod
Culture source (link if possible):  Thoughtful gift from Dr. Andrew Rhyne
If algae, CCMP # (Optional): 
http://ccmp.bigelow.edu/
Culture Establishment Date:  1/8/2013
Continuation Date:  4/8/2013

Culturing Vessel Details
Salinity:  1.020 - 1.026
Temperature:    72-74 F
pH:  Not measured

Vessel description:  Various, but mostly 2L borosilicate media jars, and 1000 mL Erlenmeyer flasks
Lighting description:  Ambient fish room light. 
Lighting cycle:  Typically 18 hours on / 8 hours off.
Aeration description:  Very light.  Rigid airline set near the bottom, and about 1-2 bubbles per second.

Methodologies
Split methodology: Various.  See notes in Journal.

Culture medium description: 
New, aged artificial salt water.  Fed live phytoplankton, usually a monodiet of Isochrysis, very sparingly and often.  See notes in Journal.

Cell count:
Varies as population of nauplii vs. adults varies over time.

Reference links:   http://bit.ly/Xxlkbe
                              http://www.sciencedirect..../pii/S0044848603001613

Additional Information
(No Pictures or Videos in the Section Please)
Notes: 



You will be required to provide photographic evidence and as much detail as possible about your project in this thread.
If your thread does not contain detailed enough photos  and information the MBI Council will not be able to approve your reports.

Here is a composite image I assembled from about 6 different images of the same individual copepod I took today with the microscope/camera combo I have access to at work.  I think this is an adult female.  It was one of the largest individuals I could find in the sample I was working with.
 

Attached Image(s)

I used the software at the lab to measure the smallest nauplius I could find (I think it may be an N1 nauplius  [EDIT:  After more reading, I'm now almost certain that it is an N2 nauplius, and not an N1], but I'm not sure) and also the largest adult I could find.

The nauplius is 45 microns wide, and 82.6 microns long.  The adult has a prosome length of appx. 400 microns, a Total Length of appx. 530 microns, and a width of appx. 200 microns.
 
Here are the images:
 


 
EDIT:  Note that the nauplius was photographed at 400x, while the adult was photographed at 100x.

Attached Image(s)

Here is a combined image of the adult and the nauplius at the same magnification factor.
 

 

Attached Image(s)

Great pics!

http://www.fishtalpropagations.com/#!home/mainPage
"Making captive breeding easier."

The N1 is tiny and more square, lacking the bud at the posterior. That bud becomes hooked shaped shortly in N3-N6.  We're getting ready go from egg to adult to egg in a control study to look at duration and size distribution.   I can get you that information. 
 
 

Great info Jim. Its funny the timing of this in contrast to the info I was looking at for the gobies. I'll scope my cultures to see what I have this weekend.

-Adam

Great photos, Jim!

--Andy, the bucket man.
"Not to know the mandolin is to argue oneself unknown...." --Clara Lanza, 1886

I have found some good information in "Handbook for the Cultivation of two Hawaiian Paracalanid Copepods" by Kyle VanderLugt and Petra H. Lenz.  The PDF can be found here:  http://bit.ly/Xxlkbe (please forgive them for getting Harpactioids and Calanoids mixed up in the caption of Figure 4 on Page 5).
 
The good stuff starts on page 12, where Section 4 on "Copepod Cultivation" starts.  There is a primer on phytoplankton production and the use of a hemacytometer to measure algal density.
 
The algal density part would seem to be an important part of maintaining healthy Parvocalanus cultures that is, in my interactions with other hobbyists, almost entirely overlooked.  I think that having a reasonable idea of the density of your algal cultures, and thus, how much algae to feed the copepod cultures, may be one key to success with Parvocalanus.  The density recommended in this Handbook ranges from as low as 5,000 cells per ml up to 100,000 cells per ml, depending on copepod density and whether high nauplii production is desired.  They also note that while it is important not to let the copepods starve, problems can arise if the algal density gets too high.
 
Isochrysis cultures near the end of the exponential phase or approaching the plateau phase of growth may contain anywhere from as little as 1 million cells per ml up to perhaps 15 million cells per ml, depending on many variables in the phytoplankton culture protocol used.  In order to know how much algae to add to your copepod culture, you have to have some idea of how dense it is in the first place.  Florida Aqua Farms sells simple Secchi algal density measuring sticks (http://florida-aqua-farms...ae-density-measurer/), together with a simple chart for estimating algal density for various species of microalgae.
 
If I am able to come up with a good approximation of how dense my algae culture is, then I will be able to intelligently add algae to my copepod culture to ensure that they are being fed enough, but not being overfed.  Terms such as "lightly tinted" or "weak tea", or other such subjective descriptions of what a given algal density is supposed to look like have always bothered me.
 
I did bring some of my Isochrysis into the laboratory where I work yesterday, and used the hemacytometer there to do a cell count.  The result were interesting.  While the Secchi stick / chart method of estimating the density indicated I should have had appx. 15 million cells per ml (around 3.5 cm on the Secchi stick), the hemacytometer count returned a result closer to 6 million cells per ml.  Granted, it was the first time I used a hemacytometer, and talking to the microbiologists at the lab, they weren't surprised at all by the discrepancy, and they said that considering the potential for sampling errors and numerous other variables, they considered my count to be relatively close to the Secchi reading.  The Secchi stick method is subject to variables, too, such as the amount of light in the environment when taking the reading.  I plan to do some more cell counting soon, though, to see if I can get a better sense of what my actual algal densities are.
 
I could keep going on, but will stop now, before this post turns into a book!
 
 

i have no problem with a book… keep going, please.
 

check out Kathy's Clowns, llc website:
http://kathysclowns.com
Captive bred clownfish and more
(Wholesale to the trade.)

Let the book continue:
 
When it comes to quantifying algal density, cells per milliliter is an obvious commonsense measurement.  The scientific literature, though, often refers to algal density in terms of units of the carbon contained within the algae per given volume, expressed as micrograms of carbon per liter, for example.  This allows for a consistent delivery of algal nutrition regardless of which species of algae, or mixture of algae, is being used, as long as you know the amount of carbon per cell of the various algae species being used as food sources.  With this approach, you establish a target level of carbon density, usually expressed as micrograms of carbon per liter in the culture media, and then determine the appropriate cell density based on the carbon content per cell of the algae being used, which is usually expressed in terms of picograms of carbon per cell.  This target level is frequently expressed in terms of a "saturation level" of carbon for the organism in question.
 
Let me give a practical example.  In the paper, "Egg production, egg hatching success and population increase of the tropical paracalanid copepod, Bestiolina similis (Calanoida: Paracalanidae) fed different microalgal diets" by Caymus, et. al., 2009, they feed the Bestiolina a "saturation level" of 1500 micrograms of Carbon per liter, which is "known to saturate copepod feeding", according to another paper, "Bioenergetics of the planktonic copepod Acartia tonsa: relation between feeding, egg production
and respiration, and composition of specific dynamic action", by Kiørboe, et. al., 1985.

I haven't yet found a good source for determining the carbon content per cell of various algae species, aside from the data that can be seen in Table 1 in the paper, "The potential of tropical paracalanid copepods as live feeds in aquaculture" by McKinnon, et. al., 2003, and I have yet to determine the source of the data in that table.  Nonetheless, according to that table, Isochrysis has 17.8 picograms of Carbon per cell. 
 
Remember that we are trying to translate a target value of micrograms of Carbon per liter into a number of cells of algae per milliliter.  That implies a factor of 1,000 (liter => milliliter).  Remember also that we are translating micrograms of carbon per liter into picograms of carbon per cell.  That implies a factor of 1,000,000 (micrograms => picograms).  The 1,000,000 divided by the 1,000 equals a factor of 1,000 (picograms per cell => micrograms per liter).
 
That said, assuming a target Carbon concentration of 1,500 micrograms of Carbon per liter, and a Carbon concentration of 17.8 picograms of carbon per cell for Isochrysis, that means that we will need 1,500 / 17.8 * 1,000 = appx. 84,270 cells per milliliter of Isochrysis to achieve our target density.
 
There are various other references to algal density for culturing Paracalanid copepods (many studies have involved Bestiolina rather than Parvocalanus, but they are closely related enough that IMHO, Bestiolina data is quite relevant to Parvocalanus culture).  Various scientific papers about either Parvocalanus and/or Bestiolina (e.g., "Intensive Cultivation of the Calanoid Copepod Bestiolina similis" VanderLugt 2005) tend to indicate that the best results, in terms of survival rates and egg and nauplius production, can be achieved with algal densities equivalent to somewhere between 10,000 and 100,000 cells per milliliter for Isochrysis.  The value arrived at in the previous paragraph is right in line with these guidelines.
 
 
 

phytoplankton cells are not a set size though, they change slightly with growing conditions, thus the carbon contentcan be different per cell.  This is why we don't offer cell counts on our Instant Algae products, it depends on what time of year it is.

I'm not convinced that the Isochrysis growing in a closet in my basement at a constant temperature is really affected by the day length outside.  Still, assuming that there is a slight variation, if one is using a turbidity / light attenuation method of estimating cell density, then it would take fewer cells of a larger size, or more cells of a smaller size to attenuate the light sufficiently to achieve a given reading on my Secchi stick.  In other words, the carbon content for a given Secchi depth should even out, even if the cell count varies, wouldn't you think?
 
What I am attempting to do in this journal is to document my thought process as I try to apply the information contained in carefully controlled scientific studies to my hobbyist implentation in a practical way.  If I get close enough to maintain my Parvocalanus cultures instead of killing them (as I have done multiple times in the past), then I will have achieved my goal.
 
Now that I have an idea of the target concentration in my copepod culture, and also an idea of how dense my Isochrysis culture is, I can proceed to estimate how much algae I need to add to my copepod culture water to establish that concentration.  For instance, if my Isochrysis has a density of 6 million cells per ml, and I want to end up with around 84,000 cells per ml in my copepod culture water, then I need to add 1,000 * 84,000 / 6,000,000 = 14 ml of algae to each liter of culture water.
 
What does that look like?  A lot like water without any algae in it, frankly!  Here is a picture two flasks, one with and one without 14 ml of Isochrysis that I estimate to be 6,000,000 cells per ml:
 

 
It reminds me a lot of trying to read a phosphate test at the very low end.  That's some pretty weak tea, and it's also VERY lightly tinted.
 

Attached Image(s)

Temp is not just what effects it, nor just photo-period.  Lighting type (spectrum, pulse rate, etc), salinity, nutrients, etc all play a roll as well.  Of course the sun won't effect it in your case, but I was not suggesting that.  Sometimes I wonder why I even post here with comments like that, seriously.

I am trying to understand how significant the effect you are describing is to the practical application of my breeding operation.  Even if there is some minor variation in the cell size due to uncontrolled variables, I would expect that variation to be slight in comparison to other variables I'm dealing with, such as the accuracy with which I can estimate the cell count in my phyto culture.  I'm just trying to get somewhere close to the right density.  When published accounts of what the "right density" is ranges from 10,000 to 100,000 cells per ml, then there is an order of magnitude or more between "too little" and "too much".  I'm just trying to land somewhere in that range.  Unless I misunderstand, and the effect you are describing can cause a really significant change in the carbon content per cell (greater than the likely measurement error of my eyeballing it with a Secchi stick, for instance) then I'm just not sure this is an effect I need to factor in.
 
I do appreciate your comments and input.  I'm just trying to process your comment in the context of what my goal is here.  I'm sorry if my flip response questioning the significance of the effect to my process offended you.

Obviously you still cannot grasp what I did not like as you are repeating it yet again . Carry on, I have no desire to continue with this, at all.

Let me try a different approach, please.
 
Please help me understand how great this variation in cell size, and thus, the carbon content is, Gresham.  I'd like to learn more about this.  Do I need to learn how to estimate the cell size / cell carbon content in addition to estimating cell density?  If so, do you have any suggestions about how I can go about doing that?

Stunning pictures Jim Welsh
 

Thank you, Peerless.
 
The scientific paper "Carbon to volume relationships for dinoflagellates, diatoms, and other protist plankton" by Menden-Deuer and Lessard, 2000, in Table 4, gives formulae for calculating Carbon content per cell from cell volume for various types of planktonic organisms.  Using the formulae in that table, and calculating the volume of Isochrysis as a prolate spheroid ("Biovolume Calculation for Pelagic and Benthic Microalgae", Hillebrand, et. al., 1999), with height (h) and diameter (d) ranging from h= 6.8 to 7.2 um and d = 3.8 to 4.2 um ("A comparative study of the inhibitory effect of diatoms on the reproductive biology of the copepod Ternora stylifera", Ianora, et. al., 1995), gives a minimum pg of Carbon per cell of 16.5, a maximum of 20.6, and a median of 18.5, which is a range of +/- 11% from the median.  This variation is far less than the accuracy of my ability to estimate cell density using a Secchi stick, IMHO.
 
Moving on....
 
Having established an initial density of algae appropriate for feeding Parvocalanus, the next question is how quickly the algae is being cleared, and thus, how much additional algae needs to be added over time to maintain the desired density.  While there are studies about clearing rates, it strikes me that one could simply monitor the turbidity / light attenuation of the culture water to determine how much additional algae needs to be added to return the density to the desired level.  Clearly, one could use a hemocytometer regularly to perform cell counts, but this is a time consuming and tedious endeavor (albeit a robust one).  If a quicker and easier method of estimating the cell density could be developed that was sufficiently accurate and precise, this would be desirable.
 
Setting aside the algal density question for a moment, another potential area of concern is the density of the Parvocalanus copepods themselves in the culture.  Studies have shown that excessive density of adult copepods can have an adverse effect on nauplii production, e.g., "Management of nauplius production in the paracalanid, Bestiolina similis (Crustacea: Copepoda): Effects of stocking densities and culture dilution", VanderLugt and Lenz, 2008.  In short, assuming that you succeed in producing a large number of copepods in your culture, and a large number of them survive to adulthood, it is very likely that the increased density will result in a significant decrease in egg production, and possibly cause a sudden crash of the copepod culture, once the adult population dies off due to the limited natural lifespan.  In other words, if you are successful at culturing these copepods, it is important to regularly split, harvest, dilute or otherwise manage the adult density in order to sustain a relatively high nauplii production rate.  More on this to follow....
 

I took another sample of my dense, dark Isochrysis that measures 3.5 cm on my Secchi stick (appx 15 million cells per ml, according to my FAF chart) into the lab today, and did a more careful and scientific hemocytometer measurement of the density.  First, I dealt with the motile nature of the cells by doing a 2:1 dilution with 70% denatured ethanol, thus killing the cells, making them settle more quickly, and much easier to count.  Second, I did two measurements, and averaged the two results.  In the first measurement, I counted the cells in all 25 squares of the hemocytometer grid.  The count was 665.  So, 665 * 10,000 * 2 (dilution factor) = 13,300,000.   In the second count, I counted the 4 corners plus the middle square, and multiplied the result by 5.  The count of the 5 squares was 166.  So, 166 * 5 * 10,000 * 2 = 16,600,000.  The average of 13,300,000 and 16,600,000 = 14,950,000, which is pretty darn close to 15,000,000!
 
I also took some pictures of my Isochrysis, and used the measuring software to get measurements of about 20 randomly selected cells.  The average width was 4.36 microns, with a minimum of 3.55 and a maximum of 4.93. The average length was 6.37 microns, with a minimum of 5.72 and a maximum of 6.87.  Based on these values, the average cell volume using the formula for a prolate spheroid is 4.36 * 4.36 * 6.37 * pi / 6 = 63.0 cubic microns.  Plugging that value into the formula described in my previous post by Menden-Deuer and Lessard 2000, the picograms of Carbon per cell = 19.7.  Using this value, the target cell density to achieve the "saturation density" of 1500 micrograms of Carbon per liter = 1500 / 19.7 * 1000 = appx. 76,000 cells per ml.  In other words, I will need to add 1000 * 76,000 / 15,000,000 = appx. 5.1 ml of my dense Isochyrsis to each liter of copepod culture media to achieve the target "saturation density".
 
I brought the hemocytometer home with me for the weekend, and hope to be able to work with such dilutions and confirm the target cell counts.  At the same time, I am going to try to work up a colorimetric method for quickly and easily estimating the algal density of the copepod culture water, but more on that later.
 
By the way, I've been observing my various cultures of Parvocalanus over the last several days.  All of them are doing well, and all of them have numerous nauplii visible, as well as numerous adults.  A general definite trend is that the cultures that are more dense in terms of adults tend to be less dense in terms of nauplii, and vice versa.  The general concept that an excessive adult density suppresses egg/nauplii production seems to have merit.

Jim, this is an excellent communication of your work. It's the kind of thing we need to understand how to manage these live foods well. Please continue! 

check out Kathy's Clowns, llc website:
http://kathysclowns.com
Captive bred clownfish and more
(Wholesale to the trade.)

The more I keep Googling different search phrases related to algal densities and the cultivation of copepods, two trends emerge.  First, while the best results seem to almost always be achieved with a mixture of different algae, the overall best performing single algae is Isochrysis galbana.  Second, the clear consensus is that an Isochrysis cell density of somewhere between 10,000 and 100,000 cells per ml is almost always used.
 
That said, I did run across an interesting paper, "Development of aquaculture technology for the flame angelfish, Centropyge loriculus" by Laidley, et. al., 2008.  Aside from having a lot of interesting stuff about breeding flame angels, it also discusses their efforts to improve their Parvocalanus culture protocols.  It says, "We are continuing to refine culture conditions having optimized feed density (~300,000 cells/ml), temperature (~25ºC), photoperiod (little effect), salinity (22ppt) and culture
density (~1/ml)...."  This paper also confirms that high densities of adult Parvocalanus results in reduced reproductive activity.
 
My idea of trying to come up with an easy colorimetric method for approximating cell density at these relatively low levels has some mixed results.
 
For those of you already familiar with the Merck D-D Phosphate test kit, please bear with me here.  I have said test kit, and I really like it.  Unfortunately, it is no longer available in the USA.  It does a very good job of detecting relatively low levels of phosphate.  Like many aquarium test kits, it colorimetric in nature.  Detecting phosphate at low enough levels to be meaningful to the reefkeeper isn't easy, but this test kit does the trick, IMHO.  It uses a pair of glass vials, each of which holds 20 ml of tank water, that rest in a black styrofoam holder with two holes drilled through it to accommodate the two vials.  Here is a picture of the kit, where you can see the vials and the styrofoam holder on the right, and also a picture of the color comparison chart:
 


 
The idea is that one vial holds untested tank water, and the other holds the water that has undergone a color change due to the test.  You then place the vials, encased in the black styrofoam, over the color chart, with the untreated water over the colorized circles, and the treated water over the plain white circles.  When the color of both vials matches when looking down through them at the color chart, you have determined the amount of phosphate.  The black styrofoam helps to avoid having the color of nearby objects interfere with the subtle color difference perception necessary to achieve the desired resolution with this test kit.
 
My idea was to see if I could re-purpose the D-D Merck vials and black styrofoam holder, while making up my own color chart, that would be able to detect phytoplankton density in the range I'm working with in the Parvocalanus culture.
 
The "mixed" results I got are as follows:  The good news is that a density of 300,000 cells/ml is very much detectable with this method, and a density of perhaps 100,000 might be, but a density of more like 75,000 is very, very difficult to detect with this equipment.  To give you an idea, here is the preliminary color chart I have come up with:
 

 
The 300,000 cells/ml sample corresponds with the 6th color sample from the left. A 100,000 cells/ml sample corresponds with somewhere between the 4th and 5th sample from the left.  The 75,000 sample is maybe the 2nd from the left, but it is VERY hard to tell.  Perhaps if I leveraged this concept, but worked with taller vials, I'd be able to get better resolution.  I'm not giving up on this idea just yet!

Attached Image(s)

Jim,
 
  Great information, and thanks for the pictures of the test kits.  I've got something to compare results now to!  Thank you.

Jim, based off of your approx 15M cells per mL, what kind of variance should you see based off of what Gresh is saying?

-Adam

I don't know, because Gresham didn't specify how great the variation might be.  Well, that and a whole lot of other things I don't know!
 
I hypothesize that as long as a measurement method that is based in Beer's Law is used, like a Secchi stick or a color/turbidity comparator such as I am suggesting, then the net approximate carbon content per liter would remain relatively constant as cell size grew or shrank, for the same species of microalgae.
 
For example, lets take the combined Secchi stick reading, hemocytometer reading, and cell measurements I noted earlier as a starting off point.  The Secchi stick and the hemocytometer both estimate 15,000,000 cells / ml, and the average cell size was 4.36 microns wide by 6.37 microns long.  As noted above, that cell size (cell volume of 63 cubic microns) gives a value of 19.7 pg of Carbon per cell, which, when multiplied by the 15,000,000 cells in my Iso culture, gives appx. 300 micrograms of carbon per ml.  Now, let's assume that the amount of variation in the cell size is +/- 20%.  For cells that are 20% greater in both dimensions, the cell volume goes up to 109 cubic microns, and the resulting Carbon per cell goes up to 31.5 pg, and for cells that are 20% smaller in both dimensions, we get 32 cubic microns and 11.1 pg.  I would not be at all surprised that, assuming the same Secchi stick reading for all the cultures (3.5 cm), the actual number of cells in the culture with the larger cells would be porportionally fewer, and the number of cells in the culture with the smaller cells would be porportionally greater.  The Beer-Lambert law almost requires this to be true.
 
Note also that the change in cell size does not affect the carbon content linearly.  Cells 20% greater in both dimensions have 1.73 times the volume, and cells 20% smaller in both dimensions have 0.51 times the volume, while the larger cells have only 1.60 times the carbon, and the smaller cells have 0.56 times the carbon.  In other words, larger cells tend to be less carbon dense, and smaller cells tend to be more carbon dense.  This carbon density almost certainly will translate into opacity, which will directly affect the Secchi reading.  In other words, again assuming a constant 3.5 cm Secchi stick reading for all the cultures, the one with the larger cells will need to have something like 15,000,000 / 1.73 = 8,670,000 cells per ml to account for the difference in volume, but then that value will need to be adjusted back up by a factor of 1.73 / 1.60 to account for the lower carbon density, for a final value of something like 9,400,000.  Smaller cell culture computation is something like 15,000,000 / 0.51 * (0.51 / 0.56) = appx. 26,800,000.
 
Now, I am probably oversimplifying the math, but if you've followed along, hopefully, you get the idea, which is this:  For a given species of algae, the Secchi stick reading can probably be seen as a reading of carbon content, regardless of variation in actual cell size and count.  At least, that is the assumption I am going to bring into the fish room with me until I get information convincing me otherwise.  Again, this all needs to be considered in the context of the fact that we are using a very inexact measurement tool, and the precision required to get the job done.
 

It has now been 25 days since I established the culture(s) of Parvo, and they are all doing very well.  My two biggest problems are having to feed them at least twice daily in order to keep the algae from clearing, and having to split them when the density gets too great.
 
I have good news / bad news on the "quantification of the amount of algae in the cultures" front.  The bad news is, I really don't think I will be able to come up with any easy color-chart based method of determining algae density at the low levels we are working with here.  The good news is that, with just a little bit of practice, it appears that it is very quick and easy to check the algal density using a hemocytometer and a microscope!  That said, I will comment that I have not yet had any success at being able to see the hemocytometer grid in order to count the algae cells when using my Celestron digital microscope.  In order to be able to successfully use the hemocytometer, I have had to use my conventional optical compound microscope, so far.

The major learning for me is that the optimal algal food concentration is so much lower than I've been thinking it should be. It makes sense, though, and also the more frequent feedings.  My rotifers have been very productive since I put a perstaltic feeding pump on them, and the marinebio-guy has a continuous system that works well for him.  When I try the parvocalanus, I'll put a peri feeding pump on them as well.

check out Kathy's Clowns, llc website:
http://kathysclowns.com
Captive bred clownfish and more
(Wholesale to the trade.)

I'm well over a month now with these cultures, and they continue to do well for me.  I am splitting them regularly, when they get "too dense", based on my casual observation.  I have yet to actually quantify this density, but I estimate it to be somewhere between 1 and 5 adults per ml.  When I split, sometimes I harvest the largest copepods, sieving the culture through an 80 micron mesh, which allows the smallest nauplii to pass through the sieve.  I then return the sieved water back into the culture, and backwash the harvested copepods into a larval tank.  Other times, it is an actual "split", where I take appx. 1/2 of the water in the culture, and transfer that into a new culture, and then double the volume of both cultures with new salt water (e.g., I split 1 x 2 liter culture into 2 x 2 liter containers, and add 1 x 1 liter of new salt water to each, to bring each culture up to a total of 2 liters).  Sometimes, rather than splitting into a second culture, I simply feed the split portion to my 210 display tank.  I have also gradually been bringing the salinity of the cultures down to 22 PPT, per Laidley, et. al., 2008, cited earlier in this topic.

One other note:  I have noticed a definite trend:  My cultures of this copepod generally tend to have either (A) a relatively large number of larger copepodites and/or adults, and relatively few nauplii and/or smaller copepodites, or (B) a relatively large number of nauplii and/or smaller copepodites, and relatively few adults and/or larger copepodites.
 
It almost seems as though each culture cycles back and forth between the (A) state and the (B) state over time.

I have to admit, you are making me awfully curious about whether I just used too much algae paste in my attempts with these. I may just have to give them another shot when I get everything back up and running this summer. (I broke all my cultures down yesterday. I was a little sad, actually, since I'd kept them going so long.)

--Andy, the bucket man.
"Not to know the mandolin is to argue oneself unknown...." --Clara Lanza, 1886

Yeah, I was thinking of trying to work with some very light paste doses with these guys, but the only algae paste I have that isn't Nanochloropsis-based is some very old, expired Shellfish Diet.  It would be fun to try SDaquarist on them, though .
 

Yeah. The Tet is awfully large and would maybe go uneaten, fouling the container, but the Iso and Pav would be a good place to start. Yeah, I think I might just add a bottle of that into the order when I re-start.

--Andy, the bucket man.
"Not to know the mandolin is to argue oneself unknown...." --Clara Lanza, 1886

Actually, the paper, "Egg production, egg hatching success and population increase of the tropical
paracalanid copepod, Bestiolina similis (Calanoida: Paracalanidae) fed different
microalgal diets" Caymus, et.al, 2009, found here: http://bit.ly/TkgYZY shows that the best results for Bestiolina similis were with a mixture of Isochrysis, Pavlova, and Tetraselmis.  I realize that Bestiolina is a little larger than Parvocalanus, but not that much larger, and they are quite closely related, so I would expect similar results with Parvocalanus.

Jim, question for you. I'm going to try and culture this. In The Use of Calanoid Copepods in Semi-Intensive, Tropical Marine Fish Larviculture by Glenn Schipp, nutricionacuicola.uanl.mx/numeros/8/6Schipp.pdf, he talked about how he raised his copepods on a mixture of Isochrysis and Rhodomonas. Do you know if the Rhodomonas is necessary for culturing Parvo?

Rhodomonas is not necessary.  I'm using a mixture of Isochrysis and Pavlova (Monochrysis), but I'm sure that Isochrysis alone would work, based on the results published by Caymus, et.al., 2009 "Egg production, egg hatching success and population increase of the tropical paracalanid copepod, Bestiolina similis (Calanoida: Paracalanidae) fed different microalgal diets" which you can read here:  http://bit.ly/TkgYZY
 

Very interesting information here Jim!  Thanks for sharing so much.  The test kit idea is brilliant, and I think you're onto something.  Taller vials is probably what you're needing to make it easier to see the differences.

Don't let fear and common sense stop you! =]

With reference to the Val. day conversation: Thanks for the paper, Jim. I'm excited to get cracking.

--Andy, the bucket man.
"Not to know the mandolin is to argue oneself unknown...." --Clara Lanza, 1886

Not to pop your bubble, Andy, but Karen Brittain sent me the following link today:
 
http://www.youtube.com/watch?v=NluJ43lUM4M
 
There are many interesting things to be learned from watching the entire video, but the #1 take home points relevant to this topic are:
 
1)  Parvocalanus crassirostris can be successfully raised for long periods of time using artificial seawater.
 
2)  Optimum salinity for P. crassirostris is 30 PPT or greater.
 
3)  Live Isochrysis worked well as a feed for P. crassirostris.  Live Chaetoceros was not a good feed for P. crassirostris.
 
4)  All manner of algae paste (Isochrysis, Tetraselmis, Nannochloropsis) were entirely unsuitable as feeds for P. crassirostris.
 
 

Dang. You are such a bubble popper. No worries. Saves me time, trouble, and money. Dang it, though.

--Andy, the bucket man.
"Not to know the mandolin is to argue oneself unknown...." --Clara Lanza, 1886

I started cultures of isochrysis today...
30 ppt.
 

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