Interesting question, David.  

Do you wish to sample and release or just sample?  If the latter I would say that the easiest way would be to cut open the abdomen through the tergites and simply aspirate the gut contents with a fine hypodermic needle.  However, if you need to release the flies there is no simple method I can think of.

Good luck, Ian

Could you please specify what are you willing to do with the blood meal? Is that to identify their origin or to identify the parasite they may harbor?

The blood is the most versatile biological material, as it allows a large number of different analyzes, ranging from genetics health and toxicological assessments. The blood has essentially two components, a liquid phase (serum or plasma) and the cells (erythrocytes or blood clot, leukocytes) which serve to separate analysis. Depending on the type of processing, these components may be kept separate or together, and stored for a wide variety of different purposes.

The choice of which of the various processing and storage techniques are used for a given blood sample which depends on the search line which is intended to contribute. In general, the most versatile choice is: (1) use a few drops of blood for the preparation of thin smears, (2) freezing of a small aliquot of whole blood and blood for the rest, (3) separating and freeze plasma and erythrocytes.

The most critical and crucial point for a successful harvest of blood is the positioning and adequate physical restraint. Some care must be taken in collecting the blood.

Coloration: Preferably the thin smears should be stained a few hours after its preparation and fixation. After a week without being stained, there will be a slight worsening in the quality of staining, and two weeks after the loss of quality is considerable. Blades that are slow to be stained become cyan-blue tints and lose the cytoplasm, impairing the quality of the analysis that can be made (both as hemoparasites differential leukocyte count). A variety of dyes that can be used to stain smears, and will be highlighted here two techniques which can be easily implemented in rehabilitation centers:

Panoptic (DIFF QUICK): It is the fastest and simple technique allows a general examination of hemoparasites and differential leukocyte count, but not produce a high quality color. Is commercially ready in the form of three solutions (A - fixator B - Orange dye C - purple colorant). 1 Submerge the blade smear in solution A for 5 seconds (or more, there is no upper time limit) 2 Immerse in solution B for 5 seconds 3 Immerse in solution C for 5 seconds 4 Wash in tap water 5. drying at room temperature.

GIEMSA: It is the most suitable technique for hemoparasites. Is commercially available as stock solution. 1 To prepare a use solution, dilute a small amount of the stock solution (variable amount depending on the manufacturer of the stock solution, dilute to be opaque and dark purple staining) 2 Place the blade of smear on a horizontal surface, and flat 3 using a pipette, cover the blood smear with the solution of using 4 Wait 15 minutes (variable time, adjust accordingly depending on the desired result and the manufacturer's recommendations) 5 Wash in tap water for 6 Drying temperature The environment  use solution should be prepared on the same day and can not be stored; since the stock solution can be stored for long periods....

URINE AND STOOL

Guano samples (urine and faeces) may be obtained from animals spontaneously or during stress caused by physical restraint or by washing sewage (not common procedure).

4.1. COPROPARASITOLOGIA

Sampling: It is preferable to collect the largest possible volume of feces (greenish-brown fraction), removing the urates (white fraction). Molecular analyzes can use very small volumes but microscopic analyzes require larger volumes (2-5g for reading). Feces should be fresher possible. Feces in the colon dead animals may also be used. Using continuous and / or pencil marker to identify: the animal ID, date of collection.

Storage only for biomolecular analysis: Transferring a sealed vial (ideal Falcon tube) and kept frozen at the lowest possible temperatures. Alternatively, the feces can be retained in absolute ethanol 1: 1, although the high cost of absolute ethanol is limiting (apart from any derail hormonal analysis). Thawing and recongelamentos affecting the quality of samples and should be avoided at all costs. Valid indefinitely.

Storage only for microscopic analysis: Use one of the following preservatives solutions in volume equal to the volume of stool (1: 1), and store at room temperature in the shade, in a tightly closed bottle (Falcon tube or similar). Never freeze. Valid for months to years.  simple Formalin (formaldehyde 1:10): Absolute formalin (10ml), clean water (90ml).  AFA solution: 1:10 formaldehyde (5ml), 1% acetic acid (5ml), 70% ethanol (90ml).  APS solution: sodium acetate (1.5g), formalin 1:10 (4 ml), absolute acetic acid (2 ml), distilled water (92.5ml). MIF  Solution: Merthiolate or mercurochrome 2% (40ml), formaldehyde (1:10 5ml), glycerin (1 ml), distilled water (50ml).  Solution iodized Ethanol: 70% ethanol, iodine (qs for coloring "port wine").

Thanks for these responses so far.  I hope to hear from others as well.

Ian, I am not going to sample and release -- just collect and kill the insect for the blood sample.  I am trying to develop this as a means of using the flies as sentinel monitors to identify potential parasite and pathogen populations infecting the animals on which the insects feed.  In my area, large tabanids such as Tabanus (horse flies) and Chrysops (deer flies).

Nabil, initially I will just look for parasites, but later other viral or bacterial pathogens, and the animal species on which the fly has been feeding.  The latter objectives will require molecular techniques.

Walter, thanks for the extensive methodology recommendations.  These will be helpful.  We routinely do Giemsa staining for fresh blood samples, but the blood from the fly gut will be in a less familiar condition, and may also contain other symbionts or parasites of the insect itself.

For others reading this stream, I will value additional suggestions and guidance.

Thanks.  Bruce

David.

I would suggest 2 methods:

1- squashing the engorged insect on a filter paper and let it dry. Then test them by molecular methods, to identify the host (source of blood meal) and screen them for the required bacteria or parasites. Obviously, this method does not give you the possibility to observe the existing parasites.

2- Since you are investigating large insects, you may consider making thin smears from their ingested blood. You can  then stain them by Giemsa. These smears can be used for parasite observation and can also be scraped and screened for blood origin/parasite and bacteria (by molecular tools).

Hope that my suggestion is useful.

best

Dissect the fly under a stereo microscope in a drop of saline or PBS. Use watchmakers forceps , pins , scalpel depending the insect you are dissecting. Remove a small drop of the gut contents to a different area of you slide that should be as free of debris (from the fly's tissues as possible) and place a cover slip over it. Strive to have Dilute blood with a monolayer or less of red cells. Importantly, use a good quality Phase contrast microscope initially at X200 and then higher magnification. You will see only relatively large parasites and bacteria which will be hard to identify in any case. So better perform all this procedure under sterile conditions so you can attempt culture. Otherwise, blot some blood on filterpaper and use any number of PCR-based approaches to identify parasite and/or blood-host DNA. 

Good luck

If you want to examine, whether any man- or animal-biting fly is a vector of 'any kind of parasite or pathogen', you should distinguish between flies, caught BEFORE or AFTER having completed a bloodmeal on a potential host. In case of hungry unfed flies, that are attracted to a blood-host - or a trap mimiking such a host - you should distinguish them into nulliparous and parous flies.

Those that are nulliparous, have not yet taken a bloodmeal and thus are not expected to have taken up any parasites yet. You can discard those nulliparous flies (=> cave: transovarial transmission of some viruses and Babesia in ticks!).

Those classed parous (=> there are keys to differentiate them from nulliparous flies) should be dissected further into head, thorax and abdomen in a drop of saline. Already under the dissecting microscope, you can recognize (motile!) nematodes (filariae) or trypanosomes etc.. Alon Warburg (see above) already made some valuable recommendations as regards the search at higher magnification.

If you want to dissect  BLOODFED flies, you shall basically get them only, when they have just completed a bloodmeal. So you know the source of the blood. If this bloodhost has parasites in the blood or skin, you could follow their uptake and development in the laboratory by maintaining the fed flies for up to 14 days. Note, that, in the case of wild-caught parous flies,  there may be paraous flies that already hosted a parasite, before coming to take a bloodmeal.

In wild-caught flies, only very few shall be carrying any parasite or pathogen. So do not be disappointed, if you find nothing on the very first day!

For the identification of the parasites and pathogens you shall need more sophisticated methods, as has beeen mentioned above.

Let me at one personal advice: It is great fun and a lot of adventure catching vector flies on human or animal baits! Be prepared to find many 'parasites', some of which are not even pathogens. I have added a figure from our field-work on the transmission of human and animal filariae in Africa (=> www.riverblindness.eu), to give an impression of waht parasites you can find in Simulium flies.

Good luck, Alfons

Hi Bruce,

Products of blood meal taken up by haemotophagic  insects can finally end up at various anatomical position depending on the period of post digestion and the stage of parasites development

So if one is interested in harvesting the microorganisms acquired , the following sites  of the insect can be examined:

1. midgut for most haematozoan parasites

2. hind gut and the Malphigian tubules for filarial worms

3. salivary glands for sporozoites of malaria parasites.

Good luck.

n.b.:

As concerns filarial parasites, those are found:

- Microfilariae in the oesophagus or midgut, immediately after the bloodmeal. They either migrate through the hemocoel to the site of further intracellular development or they die and disintegrate within short time.

- Developing stages L1 (=> sausage forms), L2 or preinfective L3 are found intracellularily in the flight muscles in the thorax (=> for most filariae developing in flies, mosquitoes or Ceratopogonidae), or in the Malpighian tubulus of fat-body in ticks, mites and fleas etc.

- Infective L3 stages can be found in the hemocoel of all body-parts (head, thorax, abdomen), but mainly in the proboscis and head.

All these stages can be seen clearly under the dissection microscope in a drop of saline, when you dissect the vectors.

Since you want to focus on American tabanidae, you might share some light into the debate as concerns the development of filariae in Malpighigan tubules of deerflies:: This was claimed to occur by some workers in 1975 for the experimental development of Loa loa in American deerflies, but in contrast, in African Chrysops, development is said to take place in the thoracic muscles.

Good luck,

Alfons

What you will need:

Bunsen burner

Blunt probe

Dissecting needles

Micro Dissecting Forceps; Cross Action; Fine Tips

Micro Dissecting Forceps; Long Straight Fine Tip

Bonn Scissors 3.5" Straight 15mm Sharp/Sharp

Scalpel handle

#11 blades

Petrie dishes, disposable plastic

Stereoscopic dissecting microscope

Low melting point tissue embedding wax

Hayes saline*

*This saline was developed for use with mosquito dissection; it should work with other flies as well

Component g/l

NaCl 9.0

CaCl2 0.2

KCl 0.2

NaHCO3 0.1

Procedure:

1. Melt paraffin, pour into Petrie dish ~.5 cm deep

2. Using burner, warm probe, use probe to melt a suitable sized well in which to partially embed and mobilize your specimen (I am assuming your specimen is immobilized using CO2, a short stint in the freezer, &c.)You will probably want to orientate your specimen with ventral surface exposed to avoid having to dissect through flight muscles.

3. When your specimen is embedded, flood the dish with sufficient saline, remove the sternites, ventral thoracic plates, and other structures as needed.

4. Expose as much of the digestive system as you require (this may be from the beginning of the esophagus to the end of the rectum)

5. Using the cross action forceps, identify, isolate and snip the digestive element you have chosen (say anterior esophagus).

6. Using the direct action forceps, isolate the end portion of the digestive element you have chosen, and snip the structure

7. On removal, you have now isolated the digestive elements which you wish to further investigate.

8. With sterile forceps, you can express the contents in your culture system of choice, or onto a slide for staining &c.

9. The aseptic level you wish to maintain throughout the procedure will be dictated by your intended use of the gut contents. Lacking details in this area, I would generally suggest maintaining a level of aseptic discipline as you would in isolating any tissue from its native source with the intent of tissue culture. It is so much easier to prevent contamination than to eliminate it.

I do hope this helps; if I can provide further elucidation, le me know.

Neil Marshall, Ph.D.