>See: Taylor, W. R. & G. C. Van Dyke. 1985. Revised procedures for

>See: Taylor, W. R. & G. C. Van Dyke. 1985. Revised procedures for

> >staining and clearing small fishes and other vertebrates for bone and

> >cartilage study. Cybium 9(2): 107-119.

>

Potthoff, T. 1984. Clearing and staining techniques. pp 35-37 in: H.G.

Moser, W.J. Richards, D.M. Cohen, M.P. Fahay, A.W. Kendall Jr. and S.L.

Richardson, eds. Ontogeny and Systematics of Fishes. American Society of

Ichthyologists and Herpetologists. Special publication No.1.

Dear John:

I've got another recipe from a Brazilian colleague for

staining young fish. Note that I never tried it sofar

but you can contact Paulo now at [e-mail] for

more informations.

Good luck!

Dominique

---------------------------------------------------------------

MODIFIED PROTOCOL FOR CLEARING AND STAINING CARTILAGES AND BONES IN

LARVAE AND JUVENILE FISHES

Step 1 - Fixation:

For larvae - use formalin solution of 4-6_._

For juveniles- use formalin solution of 10_. _

Transfer specimens to 70_ EtOH after 2-3 days, do not keep_

them in formalin any longer.

* OBSERVATION: If otolith analysis is planned, do not use

formalin to fix the specimens. Even buffered formalin will

eventually corrode the otoliths. Preserve the specimens

with 95_ ethanol, use 5-10 parts of solution per fish volume. _

Change the EtOH after a week to ensure dehydration. Store

specimens in 70_ EtOH. Specimens freshly preserved in ethanol_

will stain as well as the ones fixed in formalin.

Step 2 - Dehydration:

Transfer specimens maintained in 70_ EtOH to 95_ EtOH for 24

hr, then transfer them to liquefied phenol solution. The

specimens will clear, and you can count myomeres and

visualize internal structures if you so desire. This step

should take six to twelve hours, depending on the size of the

specimens. Phenol is a hydrophobic alcohol and will remove

all the water in the specimen, which will enhance the fixation

of the alcian blue stain on the cartilages. Alcian blue is a

hydrophobic compound as well. If you want to revert the

specimen back to alcohol, just take it out of the phenol and

put it back to 95_ EtOH for several hours to extract the_

phenol then transfer the specimen back to EtOH 70_. Phenol_

will not corrode or destroy your specimen. You may repeat this

procedure as many times as you want. This is an old trick

that I learned from parasitologists that use this technique to

prepare worms for internal analysis....

Safety procedures:

Because phenol is a highly volatile alcohol and may produce

severe burns if it comes into contact with skin, always use

gloves while handling this product. Also, use only pyrex or

other glass containers, phenol will corrode plastic. To avoid

the volatile fumes, always maintain the containers with a lid

or a cover. I recommend using phenol under a hood, or in a

well ventilated area, othewise you may get some headache.

Step 3 - Cartilage staining

Transfer specimens from phenol directly to the staining

solution described below:

For 100 ml solution:

70 ml absolute Ethanol

30 ml Glacial Acetic acid

30 mg alcian blue powder

Maintain specimens in this solution for 24 hr, or until

cartilage appears blue. ** Do not Exceed 2 days. **

* OBS: Phenol will remove alcian blue staining, do not place

specimens back into phenol, if you want to retain the color.

Step 4- Neutralization

Remove specimens from alcian blue solution and place them in

a saturated sodium borate solution. Use 5-10 parts of solution

per fish volume. Leave specimens in this solution for a day.

Step 5 - Trypsin digestion

For 100 ml solution:

65 ml distilled water

35 ml saturated borate solution

1 g trypsin powder

Prepare the solution one day in advance, mix all ingredients

in an Erlenmeyer flask and homogenize it for about 20 min.

Store it in refrigerator, covering the top.

Remove specimens from the borate solution and place them in a

dish (custard cup or crystallizer) that has been rinsed well

with distilled water, add trypsin solution to the same ratio

of 5-10 to 1. Cover top with a petri dish. Maintain specimens

in this solution until transparent. Do not over digest, the

specimen should not get to soft. A helpful hint is to place

the specimen under a stereoscope and see if the vertebrae are

visible, if you can see them through the muscle, remove the

specimen from the digestion solution and place it distilled

water. The specimen should be transparent but not totally soft.

Step 6 Bleaching:

This step is used on specimens that have pigments.

100 ml solution

85ml 1_ KOH sol._

15ml 3_ H2O2 sol._

Transfer specimens from dist. water to the bleaching solution.

After about an Hour, you should see bubbles forming around the

specimen. Remove bubbles to ensure that the specimen does not

float, and stays immersed as much as possible. When pigment

have been removed, place specimens back into distilled water

and extract all air bubbles.

* OBS: Step 5 and 6 are normally in the reverse order in most

of the methodologies published, I have found out that

digesting the specimens before bleaching produces better

results, and one can see how far the bleaching has been

successful in the internal parts of the body. Also, digestion

is faster, and only requires from one to two days, instead of

four or five if you bleach the specimens first.

Step 7 Staining bone:

Solution:

1_ KOH solution with alizarin red._

Add alizarin to the KOH sol. until you get a dark red color,

similar to a Cabernet Sauvignon from Bourdeau...

Immerse the specimens in this solution for about six to eight

hours, transfer them to distilled water and observe the

fixation of the pigment to the bone. Target for a deep red

color. Put them back to the staining solution if you want

them darker.

Step 8 Destaining

Remove specimens from staining solution and place them in

distilled water, add a couple mls of 1_ KOH. Let it sit_

until all the stain has been removed from the digested muscle.

At this point you should be able to see all the skeletal

features. If the specimen is not clear enough, take it back to

the trypsin solution and digest it a little further. Once the

specimen is clear, place it in a KOH solution for one day and

start the preservation.

Step 9 Preservation

Preservation is a three step procedure:

1- Remove specimen from the KOH solution and put it into a

solution of 30_ glycerin and 70_ of 1

2- Transfer specimen from solution 1 to a 60_ glycerin and 40_

of 1_ KOH, for another week._

3- Transfer specimen from solution 2 to 100_ glycerin, and add_

a couple crystals of Thymol to avoid fungal growth. Store them

preferentially in vials that seal well.

During the preservation procedure the KOH and the glycerin

will finish the clarification and you should get a nice and

clear sample.

* OBS: Store your specimens in the dark, staining may fade if

the specimens are exposed to sun light for long periods.

Final remarks:

The modifications to the standard methodology have not been

published yet, I am in the process of puting together a short

note to be submitted to Copeia. Feel free to use this method

and let me know how it worked with your specimens.

If you have any questions send me an e-mail and I'll be happy

to answer it.

Have fun.

John:

I think you should skip that recommendation... The staining solution only includes 0.5-1% Potassium hydroxide (KOH) and a small amount of Alizarin Red S. Glacial acetic acid is used mixed with 95-99% ethanol to dissolve Alcian blue for staining cartilage. Cartilage staining (acid process) is done before clearing and bone staining (alkaline process).

Hydrogen peroxide (not chloral hydrate) is sometimes used to bleach

specimens before processing them. Borax is used as a buffer for the trypsin

solution.

The following is the current standard reference for counterstaining small

vertebrates:

TAYLOR, W.R. & G.C. VAN DYKE. 1985. Revised procedures for staining and

clearing small fishes and other vertebrates for bone and cartilage study.

Cybium 9:107-119.

and this one is also commonly cited.

DINGERKUS, G. & L.D. UHLER. 1977. Enzyme clearing of alcian blue stained

whole small vertebrates for demonstration of cartilage. Stain Technol. 52:

229-232.

Regards,

Sven

==================================================================

Sven O Kullander http://staff.nrm.se/~ve-sven/

Senior Curator - Ichthyology Tel [phone]

Swedish Museum of Natural History Fax [phone]

POB 50007 S-104 05 Stockholm SWEDEN e-mail [e-mail]

John,

Your message sounded quite familiar to me. I had the same problem a

few years ago when I tried to get some chloral hydrate to use in a

clearing and staining ichthyology lab. The Campus Police and DEA were

waiting at my door one morning after I tried to order it. It took some

time to convince them of my ignorance. Anyway, I was told that the

Chloral hydrate is an anesthetic used on horses and large mammals. For

staining, I was told that it serves as a bacteria and fungal suppressor.

Evidently, the combination of Glacial Acetic and Glycerin creates a

fairly rich growth media for microbes. I can't tell you the chemistry

involved, that's just what I was told. Since I couldn't purchase the

Chloral hydrate, I went ahead and just skipped it and never had any

problems. I've heard mention of substituting Sodium Azide, but haven't

tried it myself. I hope this helps out.

Bradley A. Young

South Dakota Cooperative Fish and Wildlife Research Unit

Department of Wildlife and Fisheries Sciences

South Dakota State University

Brookings, SD 57007

Phone: [phone]

Fax: [phone]

e-mail: [e-mail]

John,

When clearing and staining I've had success with:

1. Remove excess skin and tissue eg. peritoneal cavity, cheek muscles etc

2. 2% (w/v) aqueous KOH containing 0.04% (w/v) alizarin red for 2 weeks. (the KOH performs clearing function and the alizarin stains bone)

3. Store in benzyl-alcohol solution (40 parts glycerol: 20 parts benzyl alcohol: 40 parts 70% alcohol)

Modified from the following relavent pubications:

Wassersug R.J. (1976) A procedure for differential staining of

cartilage and bone in in whole formalin fixed vertebrates. Stain

Technology 51 (2) 131-133

Taylor W.r. and Van Dyke G.C (1985) Revised procedures for staining

and clearing small fishes and other vertebrates for bone and

cartilage study. Cybium 9(2): 107-119

Selby P.B (1987) A rapid method for preparing high quality alizarin

stained skeletons of adult mice. Stain technology 62(3) 143-146

Hope this helps

Brendan Ebner

[e-mail]

Lower Basin Laboratory

MDFRC

100 Seventh Street

PO Box 3428

Mildura, Vic 3502

Tel: [phone]

Fax: [phone]

Дата добавления: 2015-09-29