From: PO2::"r_001f5_sc@SOUTHAMPTON-INSTITUTE.AC.UK" 13-MAR-1996 10:59:51.92 To: Multiple recipients of list ALGAE-L CC: Subj: Macroalgae pigment extraction method. Dear Algae-L. Participants, I am looking for an effective and accurate method for macroalgae pigment extraction suitable for spectroscopic or / and chromatographic analyses. Although I have already found a lot of information on the subject, hence I am aware of methods commonly used, I would welcome any comments on the following topics, e.g. unusual problems that have been encountered, a way to improve the extraction, limit the degradation or extend the conservation if storage. - DMSO versus Acetone - DMSO versus sec-Butanol - Other organic solvents - the use of additives (CaCO3, MgCO3) - temperature, duration, light conditions - dry tissues versus fresh tissues - responses of Ulva lactuca, Ulva rigida, Ascophyllum nodosum, Fucus spiralis, Fucus ceranoides, Porphyra and Polysiphonia. Sandrine-C. Charrier Maritime Faculty, Southampton Institute, East Park Terrace, Southampton, UK. From: PO3::"fnpvt@AURORA.ALASKA.EDU" "Dr. Peter V. Tamelen" 13-MAR-1996 14:22:15.73 To: Multiple recipients of list ALGAE-L CC: Subj: Re: pesky invertebrates On Tue, 12 Mar 1996, Kopczak, Chuck BioAdmin wrote: > Would anyone out there have any thoughts on the effective removal of > invertebrate grazers from macroalgae? A grad student here is interested in > their impact on productivity and is considering some lab experiments to > quantify the effect. He is considering a variety of approaches including > chemical means to remove them. I told him I would check the list for any > ideas for potentially useful compounds. > > Chuck Kopczak > UCLA Dept. of Biology > 310-206-8196 > Researchers have used many methods to exclude grazers from macroalgae and the method used depends greatly on the type of herbivore in question. Many molluscan grazers (but not all) are deterred by copper ions, so some removal techniques include using copper based paint or copper strips glued to the substrate. These copper barriers must be wide enough so that the limpets or snails cannot "step" over them, since I have heard reports that they can do this! Mechanical manipulations also work fairly well. The old stainless steel cage is quite effective as long as there is a good seal to the substrate. More recently, Vexar mesh has been used with some success. Other researchers have used carpet or astroturf glued or screwed onto the substrate. The advantage of not using cages is that shading effects are minimized. If you are trying to manipulate crustacean grazers (amphipods, isopods, etc.), I have only heard of people using cages (either stainless steel or Vexar) to acheive this. But if you are doing your work in the lab, it seems that you could just pick all of the herbivores off and then put the alga in the tank without herbivores. An easy and somewhat effective way to remove amphipods from densely branched seaweeds may be to rinse the thallus in freshwater and all of the crustaceans are supposed to swim away. As you can see the methods are quite variable and epend on both the seaweed and the herbivore. Whichever method is used, however, it is extremely important to control for the effect of the herbivore exclusion itself. For example, if you were to use a chemical to deter herbivores in an enclosed tank, how do you know that any "herbivore" effect is due to herbivores and not due to the chemical? This same reasoning can be applied to any of the techniques mentioned above, but some techniques are easier to set up controls for. The copper barriers can be made discontinous so that herbivores have access corridors. Holes can be left in cages. In both of these cases the seaweed is subjected to a treatment with both exclusion device AND the herbivores as well as treatments which exclude herbivores. If you would like more information or have further questions feel free to contact me. Peter van Tamelen University of Alaska JCSFOS 11120 Glacier Highway Juneau, AK 99801 (907) 465-6557 FNPVT@aurora.alaska.edu From: PO3::"mgraham@SDCC14.UCSD.EDU" "Michael Graham" 13-MAR-1996 17:11:07.50 To: Multiple recipients of list ALGAE-L CC: Subj: fixing phtyo's Hello all, Does anyone have information about methods for fixing phytoplankton that do not affect their absorbance and fluorescence. I am interested in fixing kelp spores (sampled in the field) prior to their germination. I assume information concerning similar techniques with phytoplankton would be helpful. Also, it is best for my purposes if the spores are filtered in the field, transported to the lab, and respended ... therefore techniques which decrease the probability that the spores will become fixed to the filter, would be most appropriate. Thanks for any help. Mike Graham _____________________________________________________________________________ Michael H. Graham Scripps Institution of Oceanography "You're gonna need a bigger boat!" Univ. of California, San Diego 0208 La Jolla, California 92093-0208 -- Chief Brody, Jaws (1976) e-mail: mgraham@ucsd.edu _____________________________________________________________________________ From: PO3::"henley@OKWAY.OKSTATE.EDU" "William Henley" 15-MAR-1996 18:45:04.28 To: Multiple recipients of list ALGAE-L CC: Subj: macroalgal pigment extraction [via LSMTP - see www.lsoft.com] I thought this response was appropriate for the entire list in this instance, concerning Sandrine Charrier's inquiry about extraction methods for pigments from macroalgae. I have found the solvent DMF (N,N-dimethylformamide; see toxicity note below) to be a huge timesaver and to yield extremely consistent quantitative results for Ulva (and probably related genera) and Laminaria (J. Phycol. 31:325-331). Surely it would work with many other species as well, except for thick/mucilaginous algae (e.g. Codium fragile). It gets virtually all of the pigments (chlorophylls and carotenoids, but not phycobilins) without need for any mechanical disruption or filtration/centrifugation. Red algae remain pink after DMF extraction, so this may be a useful way to separate phycobilins. I have used it successfully with both fresh, fresh-frozen (ideally frozen quickly in liquid nitrogen or -80C freezer) and, at least for Ulva, freeze-dried and subsequently rehydrated tissue. Full extraction takes up to a day at room temperature (extraction efficiency goes way down at low temperatures, perhaps because the DMF gets viscous). Just drop in your samples and come back the next day! The only problem I have noticed is that if fresh tissue is allowed to dry out slightly, patches of the tissue may not extract completely; one simple solution that often works is to remove the partially extracted tissue, place it in a few ml of distilled water for 1-2 minutes, then resubmerge it in DMF. The best bet is to make sure the tissue is fully hydrated before extraction. Freeze-dried tissue, convenient because it allows pigments to be normalized to dry weight measured on the same sample, must be briefly (1-2 minutes) rehydrated first. Pigments, particularly chlorophylls, are stable for weeks in DMF if extracts are placed in a refrigerator or freezer after extraction is complete (even at room temperature stability is much higher than in acetone). I was able to get crude phase-separation of the three pigment classes in Ulva (carotenes, chlorophylls, xanthophylls) between petroleum ether (or hexanes) and 80% or 100% DMF (Mar. Biol. 103:267). Although I never pursued it, an Ulva DMF extract gave 'normal' HPLC peaks in a typical solvent system. The solvent is increasingly being used with higher plants and I occasionally see its use with algae. Dichromatic equations have been published* for Chl a and b; it would be nice if someone with appropriate capability would do it for chl c, although based on the close similarity of extinction coefficients for chl a/b in acetone vs. DMF, it is probably not too far off to simply use the acetone equations with DMF. One curious casual observation is that the Fo/Fa fluorescence ratio of chl a in DMF is much lower than in acetone. I would be remiss in my duties as our departmental safety liaison if I did not point out the considerable hepatotoxicity of DMF, and that apparently the safe exposure limit is below the odor threshold (OT = 100 ppm vs. 20 ppm for acetone). However, good lab safety practices such as wearing gloves and goggles and using a fume hood should be adequate precaution. Since DMF evaporates much slower than acetone (but vapors are very dense) and the extraction does not require grinding or excessive handling, the chances of exposure are greatly reduced. As a wise TV police sergeant used to say, 'Hey - be careful out there!' *Porra, R.J. et al. 1989. Determination of accurate extinction coefficients and simultaneous equations for assaying chlorophylls a and b extracted with four different solvents: verification of the concentration of chlorophyll standards by atomic absorption spectroscopy. Biochim. Biophys. Acta 975:384-394. Bill Henley (henley@okway.okstate.edu) Dept. of Botany, Oklahoma State Univ. Stillwater, OK 74078-3013 USA +405-744-5956 (FAX -7074) _______________ Original Message _____________________________ Dear Algae-L. Participants, I am looking for an effective and accurate method for macroalgae pigment extraction suitable for spectroscopic or / and chromatographic analyses. Although I have already found a lot of information on the subject, hence I am aware of methods commonly used, I would welcome any comments on the following topics, e.g. unusual problems that have been encountered, a way to improve the extraction, limit the degradation or extend the conservation if storage. - DMSO versus Acetone - DMSO versus sec-Butanol - Other organic solvents - the use of additives (CaCO3, MgCO3) - temperature, duration, light conditions - dry tissues versus fresh tissues - responses of Ulva lactuca, Ulva rigida, Ascophyllum nodosum, Fucus spiralis, F ucus ceranoides, Porphyra and Polysiphonia. Sandrine-C. Charrier Maritime Faculty, Southampton Institute, East Park Terrace, Southampton, UK.