The Museum of Southwestern Biology

Division of Parasitology

Procedures and Protocols

Updated: February 6th , 2014

Curator: Dr. Sam Loker Collections Manager: Dr. Sara Brant

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Table of Contents:

I. Labeling Recording Data II. Processing and Preserving Host and Parasite

Specimens Processing Vertebrate Specimens ……….. Preserving Parasite Specimens …………… Parasite Isolation Methods ………………….

Decanting Methods ……....................Fecal Floatation’s …………………….Blood Smears ………………………….

Disposal and Decontamination ………………III. Staining Procedures IV. Entering Data Into Arctos

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I. Labeling and Recording Data

Recording Data in the Field

If you are collecting host specimens in the field it is necessary to carry a field notebook to record data for each host specimen collected. Use the following format:

Day-Month-Year Country, State, County, Specific Locality Details, Latitude/ Longitude, Elevation

Collector’s number (each host specimen should have a unique number, i.e. ETG200)Host Species and Sex All other host data if relevant

Detail each organ you have examined, what was found in each organ and how you found it and how parasite specimens have been stored/preserved. Be sure to include what organs were NOT examined.

Make sure to list the associated parasite numbers (i.e. W745 or ETG212) as well as the MSB numbers ( i.e. MSBP2FLM)

Example: 25 March 2004

USA, New Mexico, Soccorro County, San Marcial Armendaris Ranch 33.7000N; 106.98725W, 5450 ft. elevation

SVB1340 Anas cyanoptera, female Liver positive for schistosomes, 5 worms preserved in 100% etoh – W346, MSBL2MNPMesenteric veins positive for schistosome, 3 worms removed, stored in same vial as worms collected from liver – W346Intestines examined for helminths – Positive for cestodes – 1 proglottid saved in 100% etoh –SVB1340 Cestode , MSBL2JKM – the rest saved in 80% - SVB1340 Cestodes, MSBL2JKN (***Note different MSB number)

No other organs were examined.

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II. Processing and Preserving Host and Parasite Specimens

Processing Vertebrate Specimens

Required Materials Large Dissection Scissors Small Dissection Scissors Vanna’s Scissors Glass petri dishes – Large and Medium sizes Large Syringe Glass or Plastic Beakers – Large and Medium Sizes Dissection Scope with Lights Squirt bottles (Water, Saline and Ethanol) Microscope slides Probes

Required Chemicals Deionized Water (DI Water) Isotonic Saline Solution ( NaCl + DI Water) Ethanol - 95%, 80%, and 70%

For frozen specimens: Remove from freezer and contain in a dissecting tray, leave the tray in the wet laboratory refrigerator for between 12-24 hours depending on size of the birds. Make sure that specimen is labeled and has all host information.

Freshly sacrificed specimens should be processed within a few hours of being sacrificed to maintain the integrity of parasite specimens. When working with live parasites take care to use isotonic saline for all examinations.

1. Contain the host specimen within a dissection tray. 2. Pinch and lift the skin directly above the cloaca/anus making a small incision

with large dissection scissors. Create an opening large enough to remove the viscera, including; the stomach, liver, kidneys, intestines and body wash. Each organ should be processed for parasites.

a. Stomachs and contents are saved as host diet data – but should be opened (especially in mammalian and fish host) and examined for helminths. Save parasite specimens as described in the “Preserving Parasite Specimens” section.

b. Livers should be kept intact and placed in large glass petri dish, using a large syringe (without the needle) perfuse the hepatic portal vein with an isotonic saline solution (DI is okay is host specimen has been

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frozen). Examine the perfused fluids first for helminths. Put the liver in a large beaker with isotonic saline. Break up the liver tissue liberating any helminths; this can be done by hand or using dissection scissors. Decant the beaker with isotonic saline (decanting methods described in the “Decantation Methods” section), isolating parasite specimens. Save parasite specimens as described in the “Preserving Parasite Specimens” section.

c. Kidneys should be removed from the host and placed in a breaker with isotonic saline. Break up the tissue liberating any helminths; this should be done by hand. Decant the beaker with isotonic saline (decanting methods described in the “Decantation Methods” section), isolating parasite specimens. Save parasite specimens as described in the “Preserving Parasite Specimens” section.

d. Intestines should be put into either a larger petri dish or a clean dissection tray, depending on their size, and spread out. Keep moist with isotonic saline. Examine the mesenteric veins under a dissection scope, checking for blood flukes. Use a probe or needle to manipulate the intestines, if a worm is found carefully extract it from the veins using Vanna’s scissors. Saving parasite specimens described in the “Preserving Parasite Specimens” section.

Once the veins have been examined, using small dissection scissors, make a continuous cut along the length of the intestines. Using a squirt bottle with isotonic saline gently wash the lining of the intestine. Then take a clean microscope slide and scrape the lining of the intestines. Keep all intestinal contents contained within the vessel. Wash the intestines with saline again, put all intestinal contents in a large beaker and decant. Decant the beaker with isotonic saline (decanting methods described in the “Decantation Methods” section), isolating parasite specimens. Save parasite specimens as described in the “Preserving Parasite Specimens” section.

e. Body Wash - Fill the host specimens body cavity with isotonic saline and then pour its contents into a large beaker. Decant the beaker with isotonic saline (decanting methods described in the “Decantation Methods” section), isolating parasite specimens. Save parasite specimens as described in the “Preserving Parasite Specimens” section.

3. Depending on the type of host specimen it may be necessary to examine other organs.

a. In mammalian hosts, remove the heart and using small dissection scissors open to examine for helminthes.

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b. In fish and raptor species, remove the eyes using dissection scissors and examine for helminths.

c. In fish look in the gill filaments and swim bladdersd. In avian species, especially water birds, examine the throat and the

tracheae. e. In amphibian and reptiles species, examine the mouth and under the

tongue for helminths. f. In reptile species examine the lungs and cloaca for helminths. g. For freshly sacrificed specimens of any species, make a blood smear

to looked to protozoan blood parasites. Procedure can be found in “Preparation of Blood Smear” section.

***Note: If the specimen has been frozen, isotonic saline can be substituted with DI water.

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Preserving Parasite Specimens

Required Materials Watch Glasses Glass Pipettes w/ Bulb Hot Plate/ Kettle Glass Storage Vials Labels and

Required Chemicals Ethanol – 95%, 80% and 70% 10% Formalin Isotonic Saline

Once parasite specimens have been removed from the host they must be preserved properly, methods may differ based on type of parasite and condition of the host specimen.

1. If the host was freshly sacrificed. First place recovered parasites in a small watch glass with isotonic saline.

Make sure that each watch glass is labeled with host information as well as the location it was recovered from (i.e. “Large Intestine”)

Once all parasites have been removed from a specific habitat within the host – remove some of the saline within the watch glass and replace with hot (but not boiling) water. This will kill and straighten/flatten the parasites.

After parasites have been heat killed they can go into their permanent storage vials. Individuals of each group of parasite (species, genus or family depending on ability to ID) goes into it’s own vial and has its own set of label. How to label specimens and recording data can be found in the “Labeling Specimens and Recoding Data” section.

Generally some individuals of each parasite type are saved in 95% Ethanol for genetic work while some are saved in ethanol between 70%-80% for staining and morphological work. Preservation fluids can vary depending on parasite type.

Special Considerations: o Acanthocephalans, need to be left in DI water overnight to allow

their proboscis to extend prior to preservation.

2. If the host specimen has been frozen. Place all recovered parasites in a watch glass with DI water. Make sure

that each watch glass is labeled with host information as well as the location it was recovered from (i.e. “Large Intestine”)

Once all parasites have been removed from a specific habitat within the host they can go into their permanent storage vials. Individuals of each

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group of parasite (species, genus or family depending on ability to ID) goes into it’s own vial and has its own set of label. How to label specimens and recording data can be found in the “Labeling Specimens and Recoding Data” section.

Generally some individuals of each parasite type are saved in 95% Ethanol for genetic work while some are saved in ethanol between 70%-80% or 10% formalin for staining and morphological work. Preservation fluids can vary depending on parasite type.

Special Considerations: o Acanthocephalans, need to be left in DI water overnight to allow

their proboscis to extend prior to preservation.

*** Notes: 1) When transferring parasites only use glass pipettes, specimens may stick to plastic pipettes. 2) Worms can be very delicate, be very cautions when moving or cutting a worm. Damage can destroy morphological features necessary for identification.

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Parasite Isolation Methods

Decanting Methods

Required Materials Large or Medium size beakers Dissection Scope

Required Chemicals Isotonic Saline DI Water

To examine organ contents under a dissection scope for parasitological exam it is helpful to isolate the parasites in a small fraction of the total fluids. This is a simple and time saving procedure for isolating Helminth parasites.

First take the organ (i.e. Liver, Kidney, ect…) and place it in a breaker with either isotonic saline (freshly sacrificed specimens) or DI water (frozen specimens). Break up the tissue; liberating the worms, this can be done either by hand or using dissection scissors. Begin decanting as follows:

I. Let the contents of the beaker settle for about 20 seconds. II. Pour off the top ¾ of the fluid and heavier material

III. Re-fill with clean fluid – and let settle for about 20 seconds. IV. Repeat step II & III until you are left with a small fraction of

relatively clear fluid that can be easily examined under a dissection scope

Fecal Floatation

Required Materials Small or Medium size beakers Compound Microscope Glass Coverslip Microscope Slide 15ml Falcon tube

Required Chemicals Floatation Solution

These methods are used to isolate microscopic parasite specimens like Helminth eggs or protozoans. The type of parasites you are able to recover depends on the

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specific gravity of solution you chose to float the fecal sample in. Examples of solutions and their associated specific gravity; Sodium nitrate (Fecasol® 1.2-1.25), Sodium nitrate saturated (1.3), 33% Zinc sulfate (1.18-1.2), Sheather's sugar solution (1.25). Procedure:

I. Take between 2 -3 grams of feces, and mix with the floatation solution in a small breaker

II. Let the sample settle for about 40 seconds, and pour off the top ¾ into a clean small beaker – you may also pass through a sieve depending on the amount of fecal material

III. Pour the filtered sample into a 15ml falcon tube, fill the tube with floatation solution to the very top of the tube – making a reverse meniscus. ***Note: The reverse meniscus is necessary to isolating parasite specimens.

IV. Place a cover slip on top of the tube and let stand for ~ 15 minutes V. Remove the cover slip and place on a microscope slide – to be

examined under a compound scope.

*** Note: See methods for collecting, preserving and disposing of fecal material.

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Blood Smears

If you are sampling a host specimen that has been dead for less than 10 minutes a blood smear should be taken to examine for protozoan parasites.

Procedure: I. Using a scalpel, nick a blood vessel and obtain a drop of blood

II. Place the drop of blood on a clean microscope slide III. Using a second clean microscope slide 1) Place the short side

directly in front on the blood droplet 2) Maintaining contact with the first slide pull the second slide back into and behind the sample 3) push the second slide along the first spreading the blood sample and creating a one cell layer thick smear

IV. Make sure slide is labeledV. Make 2 slides per host specimen

VI. Air dry as quickly as possible – you can gently blow on the slide

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VII. Once dry place in a coplin jar with Methanol to fix the smear – for 2 – 5 minutes

VIII. Store in a slide box until staining – See “Staining Blood Smears”

*** Note: In avian host’s blood should be taken from nicking the tibio-tarsi, in mammals, reptiles and amphibians it is easiest to nick the base the tail or the femoral artery.

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Disposal and Decontamination

Host Specimens

If host samples are to be prepared by their associated division (i.e. MSB Birds), specimens must be put into a plastic bag with a form including all host and parasite information and records. These specimens are then released to their respective MSB division. If host samples are not to be saved for specimen preparation they should be placed into a labeled bioharzardous bag and kept in the wet laboratory freezer until incineration.

Tissue Disposal

All waste including tissues, blood and feces that are not saved by an MSB division must be immediately discarded and decontaminated. All tissues, feces and materials contaminated with blood and/or feces must be put in a bioharzardous waste bucket. Blood in small quantities can be washed down the sink or put into the biohazardous waste bucket. Biohazard bags are to be changed and disposed of frequently.

Disposal of Sharps

All scalpel blades, needles, and broken glass (including; microscope slides and cover slips) that have been used to processes a specimen must be put in to bioharzardous sharps container. There are several of these containers throughout the wet lab area.

Decontamination of Work Area and Tools

All tools, dishes and field equipment are to be decontaminated with bleach. A 10% bleach bucket should be prepared for decontamination – instruments should be soaked in the bleach solution, then washed, dried, and put away. Use DI water when washing instruments. All tools, dishes and other equipment are to be stored in their appropriately labeled cabinet or drawer.

All bench tops are to be cleaned with either 10% bleach or Lysol and a sponge after the processing of each host specimen. Additionally the microscope knobs, drawer handles and anything touched or contaminated during specimen processing, that cannot be soaked in bleach, must be decontaminated using Lysol.

Storage of Chemicals

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All chemicals used in the processing of specimens must go back to their respective cabinets and are not to be kept on the bench top – this includes all concentrations of ethanol, formalin, glacial acetic acid and methanol. Chemical bottles that were handled during the processing specimens must be decontaminated prior to being put away. This excludes DI water and isotonic saline.

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