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Volume IV - 4.3 Basic Techniques

DFS Pyramid Logo

Orientation and Training

Food and Drug Administration

DOCUMENT NO.:

IV-04
VERSION NO.: 1.5

Section 4 - Microanalytical and Filth Analysis

EFFECTIVE DATE: 10/01/2003REVISED: 02/14/2013

The next stage of the training program is the dissection of insect specimens in order to learn insect morphology and recognize insect fragments. Practice is needed for the development of dissection skills, and the following guidance will enable the trainee to begin properly.

 4.3.1 Dissection Equipment

Dissection equipment is described as follows:

  • Dissecting microscope. Dissecting widefield microscopes are the best choice.
  • Probes. Standard dissecting probes, either straight or bent tip, are purchasable from most biological supply companies and are acceptable for general use. Microprobes are probes made by inserting a minuten or #2-3 insect pin into the end of a thin wood dowel, such as a cotton swab stick. A spatula-type specimen lifter may be fashioned by flattening the tip of a microprobe, and a micro-scapula can be fashioned from a flattened # 2 pin which is then sharpened on a wet stone or fine grinding wheel.
  • Forceps. In addition to tweezers, a pair of fine-tipped jeweler's forceps (Dumont #3 or 5, or equivalent) are needed for handling small objects, and a pair of coverslip forceps. The tips of the jeweler's forceps should be protected against damage when not in use by a sleeve of small diameter rubber or plastic tubing. Always keep a spare pair on hand.
  • Additional equipment. Insect pins, a fine (#11) scalpel, and a pair of fine tip surgical or iris scissors will be useful. Small hotplate (for boiling solutions), an alcohol lamp (if open flames are allowed), and a slide warming plate (50° C.). Disposable petri dishes of various sizes (100X10, 100X15, and tight fitting lid 65X10) are needed and lined (S&S # 8 ruled filter paper) and unlined filter papers are needed. 
  • Glassware. A wax bottom dissecting dish is useful as an arena (an alcohol insoluble pinning board) for the dissections. To make one, melt paraffin in the bottom of a small glass petri dish to the depth of approximately one-half of the height of the dish, and cool. After a period of use, the paraffin surface may become rough and full of holes. Simply remelt the paraffin and cool again to obtain a smooth surface.
  • Other useful items. Syracuse watch glasses (2 - 5/8" diameter), Coors porcelain casseroles (size 00) or 10-25 ml beakers with glass watch glass covers for operations that call for heating specimens in liquid, and various shallow watch glasses for use as lids. 
     

Note: Clean white beach sand can be wetted in a petri dish and used as a formable orientation dish for wet specimens (to orient specimens for different views under the stereomicroscope).

 4.3.2 Dissection Techniques

Most dissections are performed in liquid (usually alcohol) to prevent the specimen from drying and to better control the movement of small pieces of the specimen. Dry dissection can be performed in shallow petri dishes with filter papers wetted with 50/50 glycerin alcohol; however the specimens should be softened by gentle boiling in pure water prior to the dissection.

Dissecting a specimen essentially dismantles it. Pulling off or teasing apart is the easiest technique and works well for most large appendages such as antennae, legs, elytra, wings, or even smaller exposed appendages such as labrum and labia (top and bottom mouth parts). The insect's body is firmly held at a point near the appendage while the appendage itself is pulled off or severed at its base using forceps and the microprobes. Leg and antennal segments may be separated in the same manner.

Main body plates or structures (sclerites) can be broken free by judicial application of pressure from a probe or forceps. Ventral abdominal plates may crack however, so it is advisable to cut them free along the sides and then separate them. Cutting the plates free, especially if only a few straight cuts are needed gives more control than breaking the plates.

Mouthparts and other small structures can usually be teased free using probes. Adhering excess muscle and visceral tissue should also be teased away. Tissue that cannot be teased free can be macerated with a caustic solution (5-10% potassium hydroxide) but the specimen is thoroughly rinsed in distilled water before mounting. Gentle heat will speed this maceration process.

Slide mounting media are described in Section 4.2.3. If a specimen is very thick, it will cause the coverslip to rest unevenly. This can be remedied by propping up the coverslip with bits of glass (broken coverslip fragments), nylon fishing line, or other material strategically placed in the medium before laying on the coverslip.

Finished slides are labeled, ringed, and then stored flat, as the specimen is not fixed to the slide surface. Remember to mount only one specimen/fragment/hair per slide.

 4.3.3 Insect Morphology

A. Objective

The trainee should review and become familiar with the "Micro-Analytical Biology Workbook for Food Sanitation Control Analysts," Volume I (1981) by Don J. Vail, Jr., FDA Atlanta Regional Office, Atlanta, GA. or materials provided by the trainer. 

An excellent, additional approach to learning insect morphology is described in the exercise below.

B. Assignment

This exercise is geared towards learning insect morphology and serves as a useful tool for later analyses.  The trainee sets up a series of petri dishes, each petri dish representing a particular insect or series of insect fragments from a group of insects.  For example, the analyst will label a dish for Tribolium confusum, and another larger dish for stored product beetle mandibles, with adults on one side and larva on the other.

The trainee will perform the following two step process:

Step One:

The trainee performs a series of dissections on various stored product beetles. These are referred to as Known Species Plates. Using the air tight lid small petri dishes with the bottom lined with glycerin/alcohol wetted filter papers, the trainee labels the lid in indelible ink with the species name and source information or lot number (one species per plate.) The trainee places 3-4 intact whole adult insects of that species on the wetted filter paper and begins their dissections, one insect at a time.  The dissection serves as a learning tool for morphological terminology and how the insect comes apart.  This information will prove useful towards understanding what the fragments look like when separated from the body. (Note: this exercise will also provide some insight into what may happen in a milling, flour making operation when the insect is crushed or broken up). Complete the dissections of a group of 2-4 identical specimens; some species may show some size variation or will call for several dissections to get intact fragments representing all body exoskeleton parts. For example, some beetles (like the rice weevil) have closed coxal cavities which prevent the analyst from getting either a complete prothoracic fragment or complete coxae without sacrificing one fragment for the other. When this segment is completed, then dissect the mature larva on the same plate, keeping the fragments segregated.

Upon completion, these plates can be used throughout one's career for additional dissections or for reference.

Step Two:

Step Two consists of a series of Fragment Plates, where the trainee uses the 100 X 10 petri dishes and labels the lids describing the morphological fragments, e.g. all stored product beetle mandibles on one plate.  Once Step One dissections are completed, the trainee will code in India ink each line of a S&S #8 ruled filter paper with numbers or the actual species name of the insects dissected. When dry the labeled filter papers are placed in the petri dishes and wetted with glycerin/alcohol. Then species by species, the trainee places the corresponding fragment on the species line with a representative number of fragments present to show size variations, or larval/adult conditions if possible.  

When the analyst completes the plates, the plates serve as a rapid identification tool for unknown species, but clearly identifiable morphological fragments. The analyst can pick the fragment up on the end of a probe, and move the fragment nest to the known fragments on the plate, comparing size and gross shape or character.  Narrowing down the final identification without having to do a slide by slide analysis reduces analytical time. 

Note: The trainer identifies the common stored product insects the trainee should work with in preparing the initial plates. The trainee can and should set the plates up for a life time of learning; the plate collections will take considerable time to develop and may never be completely done throughout the employee's career.

The object of this section is simply to set up the system in which the trainee can systematically learn insects. It is difficult and tedious work, and has a very steep learning curve relying heavily on memorization skills and the ability to perform minute dissections under the stereomicroscope. The initial training time should be limited to one or two weeks to learn the approach alone, with additional time granted as needed. The time for these exercises needs to be granted to the employee as continuing education and quality assurance throughout their career.  For initial training, the trainees should not attempt to memorize the fragment to species as Section 4.3.4 covers this aspect. However, if needed, and if the trainee already possesses a strong entomological background, these two sections can be combined and taught as one section. If done effectively, the trainer/trainee may also spend some time learning more about the particular insect's biology. Descriptive literature accompanies each dissection.

 4.3.4 Fragment Recognition

A. Objective

The purpose of this exercise is to learn how to distinguish microscopic fragments of insects, and how to segregate and identify them from other plant tissues on the plate.

B. Discussion

In order to accurately distinguish microscopic fragments, the analyst needs a thorough knowledge of insect morphology, and access to a reference collection of authenticated fragments. This knowledge is a prerequisite (see Chapter 16 of the AOAC) before completion of any filth analysis.

C. Assignment

The previous insect morphology exercises (Section 4.3.3) were the starting point for learning and accumulating reference material. The learning process is never truly finished; the analyst should never pass up a new dissection opportunity in order to accumulate further knowledge and additional reference material.

There are certain qualities peculiar to cuticular fragments of insects that serve as proof of insect origin. Even though a suspect fragment may have a genuine insect appearance, it cannot be reported as an insect fragment unless there is proof of insect origin. As outlined in the AOAC, the following diagnostic characteristics are the proofs by which a positive identification of insect fragments can be accomplished.

Diagnostic Characteristics of Insect Fragments

  • Shape. The shape of a fragment is diagnostic if it is recognizable as an entire or particular portion of, an appendage, body segment, or specialized structure of the insect body.
  • Setae. The presence of one or more non-cellular setae with associated setal pit (papilla) is diagnostic. Should the setae have become separated from the fragment, the presence of setal pits is sufficient for identification. 
  • Sculpture. Surface pattern (sculpture) that is typical of a particular part of a insect is diagnostic, quite often to the family or genus level.
  • Sutures. Fragments that actually consist of portions of interlocking plates (sclerites) are, by the complex form of the joining interface (suture), proof of insect origin.

Certain qualities of insect cuticle may alert the analyst to look closely for one of the above diagnostic characters. Even though these secondary qualities are not unique to insect fragments, they are useful to the analyst in that their absence casts doubt on the insect origin of a fragment.

Secondary Characteristics of Insect Fragments

  • Texture. This is thinness combined with flexibility or toughness.
  • Luster. Insect cuticle often has a distinctive sheen that the trainee soon comes to recognize.
  • Lack of cellularity. This negative aspect separates insect fragments from many types of plant material. Some types of sculpture may give a superficial impression of cellularity, but close examination finds that plant cells exhibit much individual variability of size and configuration while insect cuticular sculpture tends toward repetitious uniformity of cell-typed units.

Comparison with authentic reference material is the final, irrefutable proof of insect origin and should be employed as often as possible. Identification of fragments to family and genus is routinely possible; occasionally species determinations are accomplished when reference materials (authentics or literature) are provided.

D. Assignment

  1. Systematically arrange the fragment reference material from section 4.3.3 so that it is most useful to the individual analyst preparing the material, OR, if the trainer prefers, integrate the reference material into the laboratory reference collection.
  2. Using the laboratory library, compile a personal bibliography of literature concerned with insect fragment recognition, especially from AOAC and FDA publications.
  3. Review and discuss with the trainer what has been learned thus far on fragment identification.
  4. Practice identifying unknown specimens supplied by the trainer until the analyst is familiar with the literature and confident in their own ability to identify insect fragments.
  5. Using any literature resources and reference material, identify at least 10 unknown fragment specimens supplied by the trainer.   Identifications are to the lowest taxonomic level supportable by literature resources and reference materials found in the laboratory.

 4.3.5 Mites

A. Objectives

The purpose of this exercise is to learn the basic morphology of mites.

B. Discussion

  1. Mite Features

    Mites are chiefly recognizable by their small size (usually 0.5mm or less), general lack of body segmentation, and four pairs of legs. In order to understand mite taxonomy, analysts learn basic mite morphology, which differs considerably from insect morphology.

    Body regions are defined in relation to the positions of the legs and mouth. Anteriorly, the gnathosoma bears the mouth and oral appendages. Following this is the propodosoma, whose area is defined by the first and second pairs of legs. Collectively, these two regions, gnathosoma and propodosoma, comprise the proterosoma. The metapodosoma bears the third and fourth pairs of legs. The remainder of the body behind the last pair of legs is called the opisthosoma, and collectively the last two regions are called the hysterosoma. The term idiosoma refers to the entire body exclusive of the gnathosoma.

    Appendages of mites are of three basic kinds, each of which may exhibit varying degrees of modification. The chelicerae are the front-most pair of oral appendages. They are basically pincer-type appendages, although in some groups they may be highly modified for specialized feeding while in other groups they may be greatly reduced. In addition to chelicerae, the gnathosoma may bear a pair of leg-like segmented appendages called pedipalps. Although generally very small, the pedipalps sometimes have the proportion of true legs, which can be distinguished by position, segmentation, and lack of claw-type structures or pretarsi. Legs are usually eight in number for adult mites, although certain immature stages may have only six legs. Like other arachnids, the mite has a six-segmented leg consisting of a proximal coxa, trochanter, femur, genu, tibia, and distal tarsus. The latter exhibits no secondary segmentation as found in insect tarsi. The tip of the tarsus bears a pretarsus that is often fleshy or membranous, and may bear one or more claw-type structures. Mite pretarsi exhibit literally hundreds of variations among the different mite groups. 

    Mite setae are basically similar in general appearance to insect setae. The base of a mite seta is slightly swollen and a papilla is usually evident. Due to a central cytoplasmic core, mite setae exhibit optical activity between crossed Nicols on a polarizing microscope. As with insect setae, mite setae may be variously modified.

    Solenidia are hair-like structures found on mite legs that differ from setae as there is no basal swelling, no optical activity, and very little, if any, modification of the basic hair-like form.

    Other mite features include a postero-ventral anus, genital structures whose position varies among species, leg and anal suckers, various sclerotized body areas called shields, and simplified respiratory structures roughly analogous to tracheae (peritremes) and spiracles (stigmata). These structures may each be modified or absent in any given group of mites.
  2. Preparing Mite Microscope Slide Mounts

    Mites are mounted on a microscope slide and observed under a compound microscope for identification. Prior clearing (SeeSection 4.2.3 Preparing Microscope Slide Mounts) may be needed. Because of its desirable optical qualities, the mounting medium of choice is Hoyer's solution or one of its variants. For general work, the specimen is mounted venter up with the gnathosoma pointed towards the bottom (south) edge of the slide. (This is so the compound microscope image will appear with the gnathosoma at top). The specimen is centered, pushed to the bottom of the drop of medium, and the legs spread as much as possible, before placing the coverslip. The weight of the coverslip may produce further leg spreading, but a coverslip that is too heavy will burst the bodies of delicate specimens. The smallest sized, lightest weight coverslip found should be used.

    Small amounts of heat may be applied to the mount to help spread the legs and aid penetration of the body by the mounting medium. Hoyer's-type solutions are not to be boiled as this affects the storage life and may release harmful fumes. (Safety note:  Work with proper ventilation to avoid breathing fumes.)
     
  3. Effects on Human Health

    The effects on human health of mites in foods have not been completely documented, but some deleterious attributes of mites are becoming evident.
    • Mites can cause considerable physical damage to stored foods.
    • Mites can impart a distinctive disagreeable, sweetish musty odor to foods they infest.
    • Some kinds of mites can induce allergic reactions, including asthma-type symptoms, in sensitive individuals.
    • Mites can transport spores of molds that will grow on and spoil food products.
    • Certain mites are potential intermediate hosts for parasitic organisms that infect mammals.

C. Assignment

  1. Read pp. 63-82 of: Gorham, J. R. (Ed.) (1981). Principles of food analysis for filth, decomposition and foreign matter ( FDA Technical Bulletin No. 1, 2nd ed.).  Gaithersburg, MD: AOAC International.
  2. Review mite slides and discuss with the trainer.
  3. Compile a list of references concerning mites found in the laboratory.  Check intra-agency documents such as the Laboratory Information Bulletins and FDA By-lines.
  4. Under the direction of an experienced trainer, practice mounting mite specimens until good quality whole mounts for microscopic examination can be produced.
  5. With the specimens mounted in "4," practice using the phase-contrast microscope.

 4.3.6 Hair Identification

A. Objective

The trainee will gain experience in the identification of mammalian hairs, especially rodent hairs, and observe the differences between mammalian hairs and feather barbules.

B. Glossary

Commensal - one who eats at the same table with others; an organism, not truly parasitic, that lives in, with or on another.

C. Discussion

The regulatory analyst is able to identify hairs or hair fragments from murine rodents or commensal rodents.  The commensal relationship is between certain murine rodents and man, and not some other commensal relationship they might have with other animals.

Murine rodents are so termed because they are placed taxonomically in the family Muridae. Rodents are all those animals placed taxonomically in the order Rodentia. Examples of rodents familiar to us include squirrels, ground squirrels, chipmunks, various field mice, cotton rats, muskrats, beavers, and porcupines. These are placed taxonomically in families other than Muridae but within the larger taxonomic unit, the Rodentia. This very general description serves us simply by pointing out that the term "rodent hair," used in general by microanalysts, is simply too general to use for describing the hair of commensal rodents. The hairs of most concern to the microanalyst are those from the commensal rodents, but not necessarily limited to these. The commensal rodents of most concern are the Norway rat (Rattus norvegicus), the roof rat (Rattus rattus) and the house mouse (Mus musculus). These animals are not native to North America but were introduced by commerce. Because of their close relationship to man and documented evidence of their role in disease transmission, they are considered probable health hazards, and evidence of contamination by these animals is weighed heavily by regulatory and health officials.

Other animals, many of them native rodents, often establish a temporary commensal relationship with man. These include various squirrels, muskrats, etc. Contamination by these animals, domestic animals, pets, and human hair are also be considered and recognized by the analyst.

It is not assumed that an analyst can learn to identify hairs by reading about their various characteristics. This ability can only be acquired by careful study of authentic specimens. Suspect material should always be compared to authentic specimens.

The basic structure of most hairs consists of an external layer of scales underlaid by a cortex of generally amorphous tissue. In the center of the hair is the central core, called the medulla. Striated hairs have discontinuous medullae that give these hairs their characteristic banded appearance. Striated hairs cause the most concern since the mammals that pose the greatest threat to world food supplies, the commercial rodents, all have striated hairs. The primary task of the trainee is to learn to identify hairs of the commensal rodents. This knowledge can then be applied to learning the identification of other types of hairs.

Hairs are examined under the compound microscope for identification. Most striated hairs contain considerable amounts of air trapped in the medulla. This air is removed by heating to prevent interference with microscopic observation by diffracting light away from the objective lens. As heating procedures vary widely, the trainee chooses a personal technique under the guidance of the trainer. The simplest techniques involve heating the hair in the mounting medium, glycerin jelly, so that the air is replaced entirely by medium (See section 4.2.3, Preparing Microscope Slide Mounts).

  1. General Microscopic Characteristics of Rodent Hairs
    • Prominent scales. Under the compound microscope the edges of rodent hairs have a serrated appearance due to the projecting tips of the external scales.
    • Clear cortex. The usual color with unfiltered light is bright hyaline green with virtually no dark spots. The cortex is also typically very thin.
    • Discontinuous medulla. This type of medulla is thought to be composed of cell remnants embedded in a solid matrix. Each cell or segment contains numerous pigment granules packed tightly in one end leaving the other end clear. An intervening clear air space separates each cell from the next. This contrasting alternation of dark pigment and clear areas in the medulla gives the hair its striated appearance.
  2. Guard Hair Characteristics

    Guard hairs are the long, coarse hairs of the rodent pelt. Microscopically, the medulla is seen to consist of several rows of segments or cells, each with pigmented and clear areas as well as separating air spaces.
  3. Fur Hair Characteristics

    The most striking feature of rodent fur hair is the zig-zag configuration of the hair itself. This is evident even at low magnification and is a result of bending of the hair at the internodes. A single row of medullary cells is typical of rodent fur hairs. These hairs are thinner than guard hairs.
    • Internodes. Fur hairs exhibit this rapid constriction of the hair diameter at one or more points along the length of the hair. The area where an internode occurs shows proportionate size reduction of the medulla.
    • Air spaces. The most singular characteristic of rodent fur hair medullae is the shape of the air space between each cell. This is typically the shape of a capital "I" with the stem of the "I" at right angles to the hair's length.
    • Cortical pegs. This small extension of cortical material into the medulla appears as a single indentation on each side of the medullary cell, usually in the pigmented area.

    A hair possessing the above characteristics is probably a rat or mouse hair and should be identified by comparison with authentic specimens. The analyst compares the sizes and configurations of all structures, including scales, cortex, air spaces, medullary cells, pigment granules, and internodes.

    Mammalian hairs can be deceptive look-alikes. Shrew hairs are virtually identical to some mouse hairs except for the tips, which are more elongated, and the scale pattern, which is asymmetrical, the scales on one side projecting more prominently than those on the other.  Squirrel and rabbit hairs are similar to rat or mouse hairs in general, but differences in the shapes of the air spaces and medullary cells can be used to differentiate them.

    Hairs of mammals are sufficiently different between families and genera to permit identification to these levels in most cases.  The analyst pursues this expertise through the study of authentic specimens with guidance from experienced analysts and from the literature.

    Note: As noted with insects and fragment identification, only time and experience, or specialized study will improve the analyst's proficiency in this area of expertise. The initial objective is to distinguish commensal rodents from non-commensal, but analysts are encouraged to continually study these materials throughout their careers through continuing education and QA programs. 

D. Assignment

  1. Read pp. 125-170 of: Gorham, J. R. (Ed.) (1981). Principles of food analysis for filth, decomposition and foreign matter ( FDA Technical Bulletin No. 1, 2nd ed.).  Gaithersburg, MD: AOAC International. 
  2. Prepare acceptable slide mounts of authentic hairs supplied by trainer.
  3. Prepare acceptable slide mounts of feather barbules supplied by the trainer and compare the barbules with mammalian hairs.
  4. Practice identifying rodent hairs until one feels confident in their own ability.
  5. Examine the following types of hairs and discuss how each can be distinguished from rat or mouse hairs:
    • rabbit
    • shrew
    • bat
    • dog
    • cat
    • human

 4.3.7 Excrement, Urine, Uric Acid (Morphological and Chemical)

A. Objective

The trainee will become familiar with the major types of animal (including insect) excrement that may be found in foods, and the potential health hazards presented, such as Hantavirus, coccidiosis, and related diseases, carried by vermin pests.

B. Discussion

Excrement is a term that may be applied to feces as well as other excretory products such as urine and various glandular substances, including sweat.

  1. Feces

    Feces is the word that is commonly used for the material ejected from the intestine through the anus. Fecal pellets are feces ejected in discrete units, as in the case of rodents and many insects. Feces consist mainly of undigested food remnants. Alternate terms are "dung" or "manure."

    Fecal pellets are identifiable by visual examination under a widefield microscope with comparison to authentic material. The salient characteristics of fecal pellets are size, shape, color, and, in the case of rodents, surface coating and embedded hairs.

    Rodent fecal pellets are elongated with pointed or tapered ends. The color ranges from tan to dark brown to black under dry conditions, and is also dependant upon what the animals were feeding. Interesting color variations may occur in rodents that have fed at bait stations, with blood inclusions observed. Immature rodents undergoing weaning may, for a short period, produce pellets of a light brown color. Color variations of these sorts are not routinely encountered alone, but are mixed with other, more typical, pellets. Size range is 5-20 mm for rats and mice, with mouse fecal pellets rarely exceeding 10 mm. When moistened, rodent fecal pellets exhibit a surface coating of grayish-white mucous. Mice do not need free water to survive, therefore typically exhibit dry pellets with heavy mucous coatings, while rats need a source of free water to survive, and have thinner mucous coating and moister pellets. To confirm the mucous coating, a small drop of water is placed on the surface, usually softening the pellet and producing a mucous like characteristic. In addition, as the animals are constantly preening themselves, embedded hairs may often be seen protruding from a rodent fecal pellet, or they may be disclosed by crushing the pellet. These hairs offer vital information and are essential in determining the kind of rodent involved, especially commensal rodents (rats or mice, as opposed to muskrats, squirrels, etc.).

    Fecal pellets of sheep, goats, or rabbits are rounded, without an intact mucous surface coating. They are usually less dense than rodent fecal pellets.

    Insect fecal pellets are generally small, although some grasshopper pellets may approach the size of rodent pellets. Pellets of the orthopterans and larval lepidopterans are characteristically barrel-shaped, having truncated ends and longitudinal ribs. Coleopterans and some other insects pass small, elongated, irregularly shaped pellets. Insect fecal pellets are often the same color as the food substrate. This is especially true of stored-product beetles; these pellets do not have a mucous surface coating.

    Due to the dietary habits and digestive processes of roaches, their fecal pellets may resemble mouse pellets in color. Generally of smaller size (1/8 inch or less), roach pellets exhibit longitudinal ridges and often have a somewhat six-sided appearance caused by pressure from the internal rectal glands prior to expulsion through the anus. Roach pellets do not have a mucous coating. Since roaches habitually eat their own cast skins, the presence of these fragments in a pellet is an additional clue to the pellet's origin.

    Caution: The contents or components of a particular pellet (mammalian or insect) should not be the sole basis for the pellet's identification but rather one of many observations, the sum total of which constitutes the basis for identification. For example, rodents living in the same environment as roaches may feed on dead roaches, resulting in rodent fecal pellets that contain roach fragments. Conversely, roaches may feed on rodent pellets, with the result that a rodent hair may occasionally be found in a roach fecal pellet. Bats are insectivores and their pellets consist almost strictly of insect fragments and exhibit no mucous coating. Therefore, the analyst carefully weighs all of the characteristics observed (size, shape, color, surface coating, and embedded components) in order to identify the source of the fecal pellet.
  2. Bird Excrement

    This term is applicable to bird droppings, which consist of a mixture of glandular excretions and feces. Bird excrement exhibits a texture varying from liquid to semi-solid. Drops of bird excrement usually take the familiar form of a chalky white amorphous material mixed with darker food and watery residues. Morphologically suspect material is chemically tested for uric acid to confirm the identification as bird excrement.
  3. Urine

    Urine is a term describing the fluid excretions of a mammalian kidney. This term also has applications, for birds and reptiles, which are not usually encountered in FDA work.

    Defilement of food with rodent urine is usually detected initially by observing urine-stained packaging. These stains exhibit a typical greenish fluorescence under long-wave ultraviolet light. Rodent urine stains often exhibit "streaking" configurations caused by the rodent urinating while running or by dragging its tail through a wet urine spot. Many times, even with dry stains, a urine odor is evident.  Suspect stains are confirmed chemically under many circumstances, as defined by agency policy.

    Safety Note: Handle these materials as biohazards; practice universal precautions.  Discuss with the trainer safety practices needed for the handling of these materials. Rodents and other animals serve as potent carriers for numerous diseases.  Diseases such as Hantavirus, Histoplasmosis, some tapeworm, and related organisms are the principle concerns.  Exhibits should be prepared for safe presentation in the courtroom, while retaining recognizable characteristics.

C. Assignment

  1. Read  pp. 201-216 of: Gorham, J. R. (Ed.) (1981). Principles of food analysis for filth, decomposition and foreign matter ( FDA Technical Bulletin No. 1, 2nd ed.).  Gaithersburg, MD: AOAC International.
  2. Examine authentic specimens of rodent and insect fecal pellets, rodent urine stains, and bird excrement.  Review and discuss with trainer.
  3. 3. Under the direction of the trainer, learn to perform the various AOAC methods for the xanthydrol chemical confirmation of mammalian urine, identify fecal materials and find the citations for confirmation of fecal material, and perform the chemical identification of bird excrement for Uric acid following AOAC method 970.13.