1/27/97

Hi!

I am going to start looking at marine bacteria grown on plates and was

wondering if someone had a good protocol for the fixation process. I will

be doing SEM so I need to get rid of the salts and maintain cell

integrity. I have used 2.5% glut in artificial sea water to fix but even

with DH20 washes, I still get a lot of salt on the specimen and the

specimen seems to be covered with a "slime" like coating. Is there a way

to prevent this? I want to look at the alignment of the bacteria. Thanks

for any help.

Phil Rutledge, Director e-mail: prutle1@gl.umbc.edu

Center for Microscopy voice: (410) 455-3582

UMBC Dept. of Biology fax: (410) 455-3875

Catonsville, MD 21228

Phil,

We routinely use 2% glut in 0.1-0.2 M sodium cacodylate. With three

washes, post-fix in OsO4, three washes, dehydration in ethanol, and CPD we

usually don't have a problem with salt precipitation. The slime could be a

mucous coat from the bacteria, therefore might need a prefixation

treatment. If your preparation includes the plate media then that could

be the problem.

A couple of references:

Watson, LP, AE McKee, and BR Merrell. 1980. Preparation of Microbiological

Specimens for Scanning Electron Microscopy. Scanning Electron Microscopy.

II: 45-56.

Krueger, DM, RG Gustafson, CM Cavanaugh. 1996. Vertical Transmission of

Chemoautotrophic Symbionts in the Bivalve Solemya velum (Bivalvia:

Protobranchia). Biological Bulletin. 190: 195-202.

Hope that helps,

Louie Kerr

Research and Education Support Coordinator

Marine Biological Laboratory

7 MBL Street

Woods Hole, MA 02543

508-289-7273

508-540-6902 (FAX)

VISIT OUR WEB SITE:

http://www.mbl.edu

lkerr@hoh.mbl.edu

We have looked at settlement of marine bacteria (and whatever else) on

glass plates, and I use my usual marine invertebrate fix:

4% glutaraldehyde in 0.1M cacodylate with 0.35M sucrose

(2.5% glut will probably do)

Wash with 0.1M cacodylate with 0.44M sucrose

Postfix with 1% OsO4 in 0.1M cacodylate (sucrose optional)

Dehydrate with 30% - 100% EtOH as usual, then CPD.

Haven't had trouble with salt sticking around.

Slime may or may not be removed - in many cases we WANTED to see the

slime (but Murphy's law dictates that it will disappear if that's what we

wanted to see). De-sliming seems to take place with thorough dehydration

with EtOH (?).

Try to minimize the amount of culture medium that gets fixed onto the

bacteria! That may be the culprit.

And remember to wear your lucky red shoes.

Tina

http://www.pbrc.hawaii.edu/bemf/microangelo

Tina (Weatherby) Carvalho tina@pbrc.hawaii.edu

Biological Electron Microscope Facility (808) 956-6251

University of Hawaii at Manoa http://www.pbrc.hawaii.edu/bem

The slime is normal. Crang & Klomparens _Artifacts in Biological

Electron Microscopy_ discusses this, and ways to avoid it.

Avoiding sea salts is different. You might try washing with a

sucrose solution adjusted to the same osmolarity as the sea water you're

using, then washing with DDH2O. If you're not having problems with osmium

precipitation after washing with sea water, you could do extended DDH2O

washes after the OsO4--osmolarity is no problem (or less of one) after the

Os.

Phil

Philip Oshel

Station A

PO Box 5037

Champaign, IL 61825-5037

(217)244-3145 days

(217)355-3145 evenings

oshel@ux1.cso.uiuc.edu

Yesterday I posted a message about the fixation of marine bacteria. I

guess I should have added that the bacteria is being grown on agar in 9cm

petri dishes. What I want to look at is the association of the bacteria

to each other without disturbing the colony. There seems to be a

particular orientation and I want to observe this by SEM without taking

the bacteria off of the medium. Any suggestion? I appreciate all of the

help so far, many thanks.

Phil Rutledge, Director e-mail: prutle1@gl.umbc.edu

Center for Microscopy voice: (410) 455-3582

UMBC Dept. of Biology fax: (410) 455-3875

Catonsville, MD 21228

You should be able to process them _in situ_ by puddling the

solutions on the colonies, exchanging them by careful pipetting. After

drying, dissect away sample areas and thin the underlaying agar. Leaving

the contents of the dishes intact in the dishes during processing should

reduce or eliminate distortion/curling during drying. This assumes drying

from HMDS. For CPD, dissect under 100% EtOH.

Philip Oshel

Station A

PO Box 5037

Champaign, IL 61825-5037

(217)244-3145 days

(217)355-3145 evenings

oshel@ux1.cso.uiuc.edu

Hi Phil. Try )s)4 vapor fixation 1-3 days (usually around 3) and then air drying. Worked recently on a couple of other samples I processed. Hope this helps.

Scott D. Whittaker 218 Carr Hall

Research Assistant Gainesville, FL 32610

University Of Florida ph 352-392-1295

ICBR EM Core Lab fax 352-846-0251

sdw@biotech.ufl.edu http://www.biotech.ufl.edu/~emcl/

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