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| Larviculture newsletter < Year 2005 < Issue 228 |
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| Larviculture newsletter < Year 2005 < Issue 228 |
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QUESTION:
I am looking for a protocol to measure pigmentation of nauplii. I am aware hatcheries usually have a visual or rating system to qualify their nauplii Q and pigmentation.
I am looking for the less "subjective" method available, possible through the use of a measuring instrument.
Q1- Is there a practical method to implement in "real time" at the hatchery level where we can use imagery (color / wave lenght) analysis, spectroscopy or what else ???? such that everyday the sampled nauplii could be measured ?
Q2- In its default, recommendations for fixations of samples, minimum sample size etc. so we could take all the samples for a pigment analysis at a latter time.
Eric Pinon
Service Aqua LLC / Seabait Ltd
2970 W 84 Street, Bay#1
Hialeah
FL 33018
USA
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COMMENTS 1:
The pigmentation is of course important in naups as indeed in PLs, and each country has its preference. A couple of decades ago, I was running a little hatchery in Thailand and learned many tricks of the trade in "coloration". The Chinese clients preferred clear animals, Thais like "rusty nails" and some clients preferred very dark animals. Consequently, we would color the animal according to client wishes. Maybe I have got the country preferences wrong, it is hard to remember, but with a little judicious addition of Marmite or Vegemite to the water column, the animals would change color in an hour or so to whatever we wished. The delighted client would cart them off and seed them in the pond or nursery and everyone was happy! The additions to the water are yeast based, rapidly consumed by the animal and indeed does more good than harm. Beware of pigmentation my friend! Stick to the old scoring standard, and on your PLs look for development, gut/muscle ratio ...etc all tried and proven.
Alec Forbes
aforbes@mfmr.gov.na
COMMENTS 2:
Many moons ago we developed a staining procedure using either Sudan black or oil red "O" (both of which stain for lipids) as a possible selection tool in nauplii stocking for larval rearing under the assumption that higher lipid levels as measured with the stains was related to nauplii quality. This involved dipping samples of nauplii in stains for a set time period and measuring ensuing colour development against a chart. Very quick, very messy and very effective, at least as far as staining lipids. Whether or not lipid level reflected anything to do with nauplii quality was never really determined; shortages of nauplii and issues in production (BVP was an issue then.... that's how long ago it was!) kept getting in the way.
David Griffith
dgriffith@seafarmsgroup.com
COMMENTS 3:
I think Alec gave you the best solution, if the color is wrong just feed them something that will make it right. Anyway, I keep inventing cures for which there are no known diseases, and one thing I've played with is color quantification using a digital camera. You can take a picture and load it on Adobe Photoshop, select the area of interest, use one of the "blur" filters to average-out the color, and use the "Eyedropper" sampling tool to get a numeric color reading. Or you can keep it simple and just save the original pictures, for eyeball-to-monitor comparison, but that's too easy and therefore unsuitable for shrimping. So you can make it more complicated as I did by writing software to "read" an image and output the average, numeric color value of the cropped image area. Some digicams can be operated from the computer, which makes things a bit simpler. Some can easily take pictures through a microscope eyepiece, and by Murphy's law it may be hard to find one that meets both conditions, but who knows, there's a huge assortment available. You may want to include a color standard in the picture and use Photoshop's color adjustments to be sure the light source or camera don't introduce unwanted hues. But then, if you need or want high accuracy as some of the digital-art printers do, things get even worse because the computer monitor needs to be calibrated; There are some expensive gadgets ("color densitometers" ?) they use for that. Preserving color in biological samples is I think an extremely difficult problem in most cases.
Julio Estrada julioe@ecutel.net
COMMENTS 4:
My previous email on this subject was only covering PLs and naups. As you are doubtless aware, in Thailand in the early 80's and I believe still today, there are/were many hatcheries that specialized only in naup production, and they in turn would sell to PL producing hatcheries who would then grow them out to PL14/15 ready for stocking in grow out or nursery ponds. It was in this domain that farmers had particular preferences on the "color" of the animals. Frankly, I think it makes little difference but stand ready to be corrected by our learned members. As has been pointed out by other forum members, the naups subsist on their yolk sacks which will influence their color. The animals from zoea onwards can be manipulated using any "colorant" such as marmite or vegemite, a vegetable base yeast extract that the animals gobble up with alacrity, thus changing their color quite dramatically. These are indeed all "tricks" of the trade and frankly I do not feel have the slightest influence on the health of the animal.
Now, if you start taking into consideration the additions of astaxanthin to the feeds, and other expensive experiments, then you should contact the manufacturers who will doubtless extol the virtues of their product. That is another story entirely and we experimented time and again with these chemical products in the Philippines and Thailand with little to show at the end of it. You certainly can manipulate color in your ready-for-harvest animals by playing with your water depth, Secchi visibility ...etc and this is probably the best way to go. It certainly is the cheapest. In the Seychelles, whilst experimenting with sub-strata in lined ponds back in the 80's, I came up with the most extraordinary color hues on my juvenile P. monodon who had adapted to the color of the sub-strata and the bloom in the ponds. Biologically, I do not think that the color has the slightest effect on the health of the animal, but it certainly is an important factor in selling the finished product. It was only two or three decades ago when the housewife did not like the look of the "stripes" on the P. monodon, that is where marketing came in!
I really cannot remember which country (nationality) fitted with each preference on the color of PLs, I do however recall vividly at my hatchery, buyers asking for "rusty nails", "dark PLs (si dum)", and clear animals "si sawang" (almost transparent).
At the same time as I was experimenting (fiddling is a better word) with color, I found that sensory characteristics (taste) presented a much more important and relevant field using additions and removals of certain amino acids, glycine, glutaic acid, leucine and proline ..etc and would strongly recommend further experimentation in this area. It has been amply documented by Sumidu and Hujita 1954, Hashimoto, 1965, Rangswamy et al 1970, Castille, F.L.Jr and Lawrence A.L. 1981 and Forbes et al 1988. I am delighted to see the interest in color, but I feel it would behoove us more to look at taste.
Alec Forbes aforbes@mfmr.gov.na
COMMENTS 5 :
It seems we are taking the direction of color/imagery. Why not ? I should now be a bit more specific in my need to focus better. As some of you know I work with a company specialized in raising marine polychaetes widely used in shrimp broodstock feeding (maturation). In effect we have been researching a lot these creatures and found mechanism to increase (between other) carotenoid pigments in the tissues of this fresh feed. "If" carotenoids are of any benefits to shrimp in their early stages (read nauplii) before initial feeding; we believe this form of pigments will be much more bio-available to the breeders and to their offsprings. This is the theory, how do we get to real life, measure it and see if it is worth to the industry? So within all the variable in shrimp maturation and its breeding, I am looking for a practical method to measure "original" nauplii pigmentation according to different level of carotenoids applied through our polychaetes. Not trying to trick here but look for the correlation between input and output basically. Lab analysis seems difficult because of the minimum size of the sample. So maybe imagery would work. Other describe me a bit more about the hardware and set-up we would need to capture the pictures ? I assume the color qualification could be done latter with all the samples at once.
Eric Pinon
epinon@serviceaqua.com
COMMENTS 6:
I understand a bit better now your question. Polychaetes (live) to your broodstock is a great diet and I am more of the opinion that your live polychaetes to your adult broodstock would contribute greatly to natural pigmentation without any additives at all. The polychaetes might also serve as a good vehicle to get pigments, medecine ...etc into your large animals, just as we use enrichend 1rtemia to get items into our larval and post larval shrimp. Forgive the pun, but it is a different kettle of fish to coloration of naups and PLs which is how this discussion began.
Alec Forbes aforbes@mfmr.gov.na
COMMENTS 7:
A straightforward color photograph of, say, a PL's HP, however manipulated, will only give you limited information, because a particular color can be achieved by several combinations of colored substances. It may come from a complex colored molecule of biological significance (chlorophyl, for instance, or one of the carotenes). It may be a substance that absorbs strongly at most wavelengths, transmitting mostly yellow, or another one which transmits mostly red and green. Either way, you will get a yellow picture. The color might be achieved by a combination of innocuous vegetable dyes, the kind used for pastry decoration. Or Doc Alec's yeasties. But if naups, PLs or your worms only accumulate a limited number of colored substances, there is some chance that this simple color information would be enough, if you periodically correlate color to more formal (and expensive) analyses.
A rewind to theory: Any particular color can be numerically described by several methods, according to how "color space" is defined. Only three numbers are needed in most cases. In computer-related applications, "RGB color space" is generally used, because monitors and graphics processor work by calculating and outputting Red/Green/Blue light in varying intensities. And the magic numbers in this case are called, simply enough, R, G and B. In current versions of Windows and software such as Photoshop (insert trademark warnings here) each of the magic numbers can have a value of 0 to 255, which gives you over 16 million colors. If R, G and B are 255, you get white, if they are zero you get black, if all three are 128 you get a neutral gray; 255-0-0 is a bright, pure red ... you get the drift. I can think of at least 3 other "color space" systems in use, but all of them tell you nothing about how the color is achieved. Except the process-color printing method, which uses four inks and 4 magic numbers ...
Spectrometry / spectrophotometry works with far more datapoints, therefore can yield more information. It will often allow you to determine what particular substance is causing the color with a fair degree of certainty; Even to quantify the concentration with some accuracy if the extraction method is carefuly standardized. There are a few ways to get a spectrum. Let's stick to "absorbion spectra". I think the currently popular way to do it is to illuminate the sample with nearly monochromatic light at a particular wavelength and measure intensity of the transmitted light as a fraction of the original light beam's intensity. Then change to a slightly different wavelength and repeat. Continue until you cover the full range of wavelengths, usually 400 to 700 nm which is the human-visible range, and you end with a graph which will show absortion peaks, and for many substances their characteristic peaks are well known. It is even possible to get a spectrum for solvent-extracted chlorophyll and determine proportion of the three chlorophyll types present.
I'm nearly sure there was a method in the "standard methods for seawater analysis" which involved measuring at only 4 or 5 wavelengths and getting quantitative chlorophyll informations, but this a special case for well-studied substances. More often you would need the full spectrum.
Now to the gadgets. If you have a lot of patience, it could theoretically be done with a standard shrimp-lab spectrophotometer, just set the wavelength to 400 nm and measure absorbance, change to 403 nm and measure again, continue until you reach 700 nm or fall asleep, whichever comes first. You might need to recalibrate via blank and standard now and then, between naps. After a night's rest you can plot out the 100 datapoints and get a spectrum. Not very promising for regular use. I know the Hach DR/2500 can be programmed to measure sequentially at several wavelengths, IF you buy the avanced software option. Not sure if just a few wavelengths, or if it will let you do a full scan. Maybe the Hach people have other models designed to do the sweep and output the data to a computer which can do the graph. Getting into grounds closer to what you might want, I did run by some work done (chlorophyll yet again) by way of a microscope-spectrometer combination. The researchers could actually look at individual spores of some phytocritter in vivo and get a spectrum. Will try to find it again, surely they mentioned the brand and model of gadgets used. Prices on these things should be coming down quite a bit, but probably not enough yet to fit my toy budget. So I have not looked them up.
To make life more interesting, there is also fluorescence spectrometry, which should allow further differentiation of substances present. I know even less about that, so cannot contribute additional significant confusion. There is paper chromatography and HPLC in case additional complications are needed. The first yields colors, so should be fun, the second you probably know much better than I.
Julio Estrada julioe@ecutel.net
COMMENTS 8:
Nauplii color does change dramatically with the pigments used on diets for your maturation animals and could be measured with photography if you manage same micro, same light, same camera.
John Birkett jbirkett42@yahoo.com
COMMENTS 9:
Although it is said that nauplii coloration is correlated to nauplii quality, I have not seen any scientific data on this. Logic thinking would indeed suggest that N quality is related to carotenoid levels, and carotenoids will affect coloration. In this case, pale offspring would be of inferior quality. What we do at INVE's test centers is to read the code on a Pantone color card (www.pantone.com) and take pictures with a digital camera for comparison with previous batches (nauplii and Pantone card next to each other on the same picture). Take the pictures always with the same camera and under the same light conditions.
You can also use staining methods (for total lipids; cfr mail David) or enzymatic-colorimetric test kits from Merck (total lipids, triglycerides, cholesterol, phospholipids, glucose) to monitor nutrient levels. The latter are described extensively in papers by Elena Palacios and Ilie Racotta (CIBNOR, Mexico), but it is not easy to link this information with N quality. Unfortunately, I do not know of a test kit for the measurement of carotenoids. You would need to extract the carotenoids with solvents and apply spectrophotometry. Of course, these are all destructive methods, using something like 1 gram wet weight or 100.000 N per sample (but pls check with above mentioned authors).
Remember that nutrient reserves are critical for the N to survive to zoea. These are maternal reserves, and the measured nutrient values or colors correlate to broodstock quality and broodstock feeding, yet I think they are independent of "tricks" like those described by Alec. Or are there tricks to color N as well? What is everybody’s feeling about the subjective visual check? I think that if they are done by experienced people and if combined with other N quality parameters, it can give useful indications.
Roeland Wouters r.wouters@inve.be
QUESTION:
We routinely use a mixture of hydrogen peroxide and a strong acid to clean our hatchery pipes. It is very effective at removing sediments and deposits. I wonder if anybody knows what the chemistry of this mixture is? Also, the sediments we get in our pipes are usually brownish black, with no foul smell, slightly oily but not too much, any idea what it might be? Organic?
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COMMENTS 1:
I bet your sediment is a biofilm, a microbial community which secretes complex sugars, proteins and whatnot to hold itself together and to form a skin which protects it from many regular disinfectants, antibiotics, etc. I think the polysaccharides are the major component of the skin, and matrix. The skin at least is usually "slimy" or "oily" while still wet. It's pretty compact inside the biofilm, so the smelly anaerobic metabolites probably don't accumulate in there. I'd guess the skin is permeable to those, else the bugs would poison themselves.
You could smear small bits out on a slide, as thin as possible, and look at it under a microscope. Wet mount at 400X might show you enough, have never tried it. You can "fix" the smear, use methanol if you have some, for a couple of minutes, or air-dry for 15 minutes or so. Stain it with whatever you have, such as dilute methylene blue. Dry and look again w/ oil-inmersion. Most of the material, 80% or more, will be the slimy matrix, so the picture won't probably be all that clear. Strong oxidizers will burn through the biofilm's skin. I'm very hazy on the chemistry of that process, to the point of NPI, and if you latch on to some good info on it pls let me know. Chlorine bleach should be as good or better than H2O2 as an oxidizer. But I wouldn't mix it with acids until someone who knows his shit says it's OK. Preferably someone who's been doing it for a while and is certifiably alive and well. Also, Ed Scura mentioned some time back that concentrated ozone does a great job on biofilms. Anyway, what you're using now is working.
Strong acids will "digest" proteins and many other organic substances, I think by breaking some of the chemical bonds, anyway that's why we have HCl in our stomach. If you're using sulphuric acid, it's also a strong oxidizer, and a dehydrant to boot. It will steal H2O away from sugars, leaving mostly carbon behind.So, the acid + oxidizer combo would act by destroying or greatly weakening the biofilm's skin & matrix, leaving the bugs naked and therefore killable and sweepable.
Mixing H2O2 mith many acids will yield some proportion of peroxy-acids, for instance peroxymonosulfuric acid: H2O2 + H2SO4 <---> H2SO5 + H2O. Some, such as peroxyacetic acid, are fairly good at digesting even refractory organics like lignin, and they're oxidants / bleachers just like H2O2. As far as I could dig up, peroxy-acids are not necessarily more effective than H2O2 itself, at least at high pH. Some of them seem to be particularly useful in removing metallic contaminants.
H2O2 is most aggresive to organics in high-pH solutions ( > pH 11.5). Perhaps too active, so it may not reach remote spots in your piping before it decomposes.
H2O2 might have a double effect on biofilms; it will weaken the slime by oxidizing its components, and the oxygen liberated once it penetrates the mess would help mechanically by literally blowing it apart. I don't know how speculative this is, didn't track down "hard" information.
H2O2 by itself will remove the "flesh" from diatom samples, leaving the silicon skeleton. For light microscopy, they're treated with HCl in a second step, but that's supposed to be mainly to remove any metals. As to details of oxidation reactions in general, never mind oxidation of organic substances, I've found it the densest going in my chemistry book once Dallas helped me with the "carbonate conspiracy". So the simplistic idea I keep in mind is that if you attack organics with enough strong oxidizers and supply oxygen, you end up disassembling the original goo to yield CO2, H2O, NO3 etc.
How about an experiment? Set up a tank so your LRT outlet water flows through it. Let some pieces of PVC piping sit in there until you get thick-enough biofilms. Then stand them in small tanks so they're partly submerged in H2O2 alone, acid alone, and your usual mixture. Keep track of how fast and how thoroughly the film is removed. You could play with concentrations, too. Should give you a feel for what's most cost-effective.
Julio Estrada julioe@ecutel.net
COMMENTS 2:
Sounds like it is a biofilm deposit. The black is probably from a small amount of Fe being converted to FeS by small amounts (less than you can smell) of H2S being generated at the base of the biofilm.I have used acid/peroxide mixtures -- about 0.1% H2O2 at a pH of < 2.5. With time, it will react with biofilms and slowly kill things. The biofilm tends to protect the bacteria, virus, etc. within the biofilm from attack from external oxidants. The organics in biofilms will react with any oxidants and neutralize them. However, the acid component will diffuse in and as long as there is no carbonate scale to neutralize the acid, the pH in the biofilm will decrease to the point where almost everything is killed (both good and bad bugs). Biofilms are a major problem in other technical areas and are very resistant to chlorine. Mechanical systems like high pressure washers (which can also be putdown pipes) are good at disrupting the biofilms and allowing attack by chemicals and detergents. Blast it with 3,000 psi water then bleach it. There are some good web sites on the subject. I have also use saturated salt tanks for killing surface biofilms. Saturated brine will suck all the moisture out of a biofilm and most of the organisms in the biofilm will die from the osmotic pressure. The killing impact is only from osmotic pressure, which will penetrate a biofilm very rapidly (seconds to minutes) without being neutralized. Makes a good quick net dip and won't harm the hands, clothing, etc. of the staff. It is very good for macro organisms like flukes, epistylis, protozoans, most aquatic bacteria, etc. No data on virus but I would imagine it would depend upon the specific virus.acid/peroxide will remove the MnOx brown stains created by using permanganate for sanitation. Permanganate is handy for adding to chlorine solutions to see if they are active. It wiill show up biofilms on surfaces by staining them (quality control checks on the staff to make sure they are properly cleaning the equipment).
Dallas Weaver deweaver@surfcity.net
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