riedi v’s ingens

Also, almost forgot…..

I am very interested in seahorse pathology. I would like to know more about using light microscopes for diagnosing seahorse diseases and preparation of samples on slides etc. I am an undergrad science student and like to tinker with microscopes.

Could you please recommend some good books on this for me?

Cheers,
Karen

Dear Karen:

A good place for you to start would be to pick up a copy of "Working Notes: A Guide to the Diseases of Seahorses." It will provide you with an excellent introduction into seahorse anatomy and has some excellent dissection and necropsy photos as well as a number of photos of seahorses with various health problems. It includes a good discussion of bacteria and endoparasites and ectoparasites, with descriptions of their key characteristics for identification purposes. It’s available online at the following web site:

Click here: Working notes: a guide to seahorse diseases > books > The Shoppe at Seahorse.org | CafePress

http://www.cafepress.com/seahorses.55655887

Any good textbook on microscopy or microbiology will include good instructions for preparing wet mounts and slides, Karen.

To get you started, here are the diagnostic procedures for seahorses as described in the latest version of the Syngnathid Husbandry Manual for zoos and public aquariums. As you will see, feces can also be obtained by performing a colorectal flush as opposed to waiting for the feces to be eliminated naturally:

<open quote>
There are several potential etiologies when an animal is experiencing buoyancy
problems. If the pouch appears asymmetrically distended or symmetrically distended with
attendant buoyancy problems, a percutaneous fine needle aspirate should be performed on the
pouch contents. Any fluid aspirated should be dried and stained with Wright-Giemsa stain, Gram
stain, and acid-fast stain. The pouch can also be flushed with sterile saline and the aspirate sent
for culture.

If a hyperinflated bladder is suspected, a bright light can be directed from behind the animal to
visualize the location and borders of the distended organ. This is useful when attempting to
deflate the bladder. The needle should be directed between the scutes/plate margins for ease of
penetration through the skin. The external area can be rinsed with sterile saline or a drop of a
triple antibiotic ophthalmic solution applied prior to needle penetration.

Diagnostic dips or baths or diagnostic washes of the branchial cavities can be performed to obtain an etiologic diagnosis since the gill tissue itself is so inaccessible. Because of the semi-closed nature of the branchial cavities, branchial washes with sterile, 0.9% NaCl are far easier and much less traumatic than traditional gill biopsies.

Skin lesions should be swabbed with a few sterile, wet (using sterile saline), cotton-tipped
applicators and evaluated by wet mount, gram stain, acid-fast stain, and/or Wright-Giemsa stain.
Skin ‘scraping’ is possible but quite difficult in practice due to the irregular surface architecture
of the bony-plated armor. Follow-up diagnostics to the initial skin swab include aerobic bacterial
culture, mycobacterial culture, and cytological exam by a pathologist familiar with fish. Fin clips
can be performed if fin lesions are observed.

Any unusual lump, bump, should be worked up with a fine needle aspirate (27 gauge needle &
TB Syringe) and sent for culture, wet mount and/or staining.

Feces can be obtained by the wait-and-watch method or a colorectal wash under MS-222 (tricaine methanesulfonate) sedation using a tuberculin syringe, 0.9% NaCl, and a 3.5 gauge red rubber catheter. Oftentimes, the seahorse will void feces reflexively after anesthetic induction with MS-222.

Blood collection is technically very difficult due to the relative absence of accessible peripheral
veins and the small size of most syngnathids. However, small amounts were obtained in a recent
study on CBCs and cortisol levels in H. erectus (P. Anderson, K. Harr and D. Heard, University
of Florida, unpublished data). They were able to successfully draw 0.2 mls from animals, even
up to 0.4 mls in some animals. Another 0.2 cc from the same two animals approximately 18 days
later. Blood was drawn at the basal part of the tail from a ventral approach. Cardiac sticks were
unsuccessful.

Radiography can be a very helpful diagnostic tool. The ideal machine for taking radiographs of
seahorses is a mammography unit in terms of optimal radiographic detail and contrast. Important
structures to evaluate include swim bladder (size, shape, presence or absence of fluid), coelomic
cavity (free air, fluid, masses), alimentary tract (aided by a small bolus of barium sulfate if
needed), liver position and size, kidney position and size, gonadal position and size, and brood
pouch contents. Larger seahorse species as well as the seadragons are particularly at risk for
foreign body (usually substrate) ingestion which can easily be ruled in or out with plain film
radiography. Ultrasonography of brood pouch contents should be made possible by using the
smaller transducers (7.5-MHz – 10.0-MHz) employed in ophthalmologic evaluations.

Sedation and Anesthesia

MS-222 at the standard fish dose of 50 – 100 ppm works quite well for most of the syngnathids.
In low-alkalinity water it is recommended to buffer the solution at a ratio of 2 parts sodium
bicarbonate:1 part tricaine (wt:wt). The seadragons have a prolonged recovery time at 100 ppm
and 50-75 ppm is the recommended dose for these two species. In a prolonged anesthetic
recovery situation, it is advisable to ventilate the animals with fresh seawater containing no MS-
222. Because of the long, rather narrow tube snout and the semi-closed nature of the branchial
cavities, assisted ventilation is easily achieved with a 3.5 to 5.0 French red rubber catheter
inserted through the tube snout to the level of the pharynx. A syringe filled with fresh saltwater is
then attached to the end of the red rubber catheter and pumped in a pulsatile manner every few
seconds until the animal is spontaneously breathing at a normal rate. The success of assisted
ventilation is easily assessed by watching the opercula move in and out. This technique has also
been successfully used to resuscitate animals in respiratory arrest. Long-term anesthetic
procedures should employ a flow-through system with oxygen supplementation in the sump or
reservoir. <Close quote>

Best of luck with your science studies, Karen!

Happy trails!
Pete Giwojna

Dear Karen:

You’re very welcome to all the information I can provide to help keep your seahorses happy and healthy, Karen!

I am in the process of updating my new book for publication right now. A lot of work still remains — I have to integrate all the illustrations into the text and then convert the manuscript into "electronically ready to print" form, but I anticipate the release of the book within the year. Of course, I will keep you guys informed as soon as it’s ready.

Happy Trails!
Pete Giwojna