[LAB DIAGNOSIS] LAB MODULE 3 UNIT 2 AND 3 Flashcards
1
Q
In surveys of infected population, as well as individual cases, it is sometimes desirable to estimate the intensity of infection by counting the number of eggs in the feces.
A
EGG COUNTING PROCEDURES
2
Q
Egg counts provide a reasonable estimate of the number of adult worms present.
A
EGG COUNTING PROCEDURES
3
Q
Egg counts before treatment may help determine whether treatment is needed and counts after treatment assess its success based on egg reduction rate (ERR) as a consequence of reduction of worm burden.
A
EGG COUNTING PROCEDURES
4
Q
This procedure uses a measured amount of stool which has been sieved
through a wire mesh and pressed under cellophane paper soaked in glycerine-
malachite green solution.
A
A. Kato-Katz method
5
Q
A uniform amount of stool is examined through the use of a template with a
uniform-sized hole in the middle.
A
A. Kato-Katz method
6
Q
A hole of 6 mm on a 1.5 mm thick template will deliver (?)
A
41.7
mg of stool
7
Q
o a hole of 9 mm on a 1.0 mm thick template, ?
A
50 mg
8
Q
o a hole of 6.5 mm on a 0.5 mm thick template, ?
A
20 mg
9
Q
All eggs seen in the whole preparation are counted.
A
A. Kato-Katz method
10
Q
The total egg count is multiplied with a factor depending on the amount of
stool used.
A
A. Kato-Katz method
11
Q
o The total number of eggs counted is multiplied by 24 for a
(1) template; by 20 for (2) template; and by 50 for
(3) template.
A
- 41.7 mg
- 50 mg
- 20 mg
12
Q
Consistency of the stool is the main determinant for the sensitivity of this
technique, since well-formed stools yield higher egg counts than moist ones.
A
A. Kato-Katz method
13
Q
The technique can only be done on fresh formed stools and not on liquid and
preserved samples.
A
A. Kato-Katz method
14
Q
The only human nematodes for which it is reasonably possible to correlate egg
production with adult worm burdens are Ascaris lumbricoides, Trichuris
trichiura, and the hookworms, all of which are soil-transmitted helminths
(STH).
A
A. Kato-Katz method
15
Q
Note: The procedure is also useful for assessing the intensity of infection with
Schistosoma.
A
A. Kato-Katz method
16
Q
Owing to its simplicity and relatively low cost, the Kato-Katz technique is
recommended by the World Health Organization (WHO) for epidemiological
surveys and surveillance pertaining to soil-transmitted helminthiasis and
intestinal schistosomiasis control programs.
A
A. Kato-Katz method
17
Q
uses a counting chamber with two compartments,
each with a grid etched onto the upper surface.
A
B. McMaster technique
18
Q
This enables a known volume of fecal suspension (2 x 0.15 ml) to be examined
microscopically.
A
B. McMaster technique
19
Q
A known weight, 2 gms, of feces and a known volume, 28 m l , of flotation fluid
are used to prepare the suspension with a total volume approximately 30 ml.
A
B. McMaster technique
20
Q
When filled with a suspension of feces in flotation fluid, much of the debris will
sink while eggs float to the surface, where they can easily be seen and those
under the grid counted.
A
B. McMaster technique
21
Q
The number of eggs per gram of feces (EPG) can be calculated by multiplying
the number of eggs under the marked areas by a simple conversion factor.
A
B. McMaster technique
22
Q
Multiply the number of eggs counted in one compartment by 100, or by 50 if
both compartments are used, to give the number of eggs per gram (EPG) of
feces.
A
B. McMaster technique
23
Q
has been developed to easily carry out the flotation of the fecal
sample (fresh or fixed) in a centrifuge, the translation of the apical portion of
the floating suspension, and the subsequent examination under the
microscope.
A
C. FLOTAC® and Mini-FLOTAC®
24
Q
These techniques use the FLOTAC® apparatus, a cylindrical-shaped device
made of polycarbonate amorphous thermoplastic, with two 5-ml flotation
chambers, which allows up to 1 g of stool to be prepared for microscopic
analysis.
A
C. FLOTAC® and Mini-FLOTAC®
25
There are two versions of the FLOTAC apparatus:
o FLOTAC-100, which permits a maximum magnification of ×100
o FLOTAC-400, which permits a maximum magnification of ×400
26
However, the main limitation of (?) is the complexity of the technique
which involves centrifugation of the sample in a specific device
FLOTAC
27
Centrifugation can take place in either a large volume centrifuge or in
benchtop centrifuge with rotor for microtiter plate, an equipment that is often
not available in some laboratories.
C. FLOTAC® and Mini-FLOTAC®
28
The (?) comprises two physical components, namely the
base, the reading disc and two accessories, the key and the microscope adaptor that are useful for assembly of apparatus and examination under a microscope.
Mini-FLOTAC® apparatus
29
There are two 1-ml flotation chambers, which are designed for optimal
examination of fecal sample suspensions in each flotation chamber (total
volume = 2 ml).
C. FLOTAC® and Mini-FLOTAC®
30
(?) be used in combination with Fill-FLOTAC® .
Mini-FLOTAC®
31
It is composed of a graduated container and a lid.
Mini-FLOTAC®
32
On the top of the lid, there are two holes with screw caps: a central one (with
a large screw cap) for the collector/homogenizer pole and one (with a small
screw cap) to which a tip is attached for passage of fecal suspension.
Mini-FLOTAC®
33
The conical collector permits measurement of 2 g of feces (Fill-FLOTAC 2).
Mini-FLOTAC®
34
The filter in the lower part of the lid has 250-µm holes to ensure an optimal
filtration of the fecal suspension.
Mini-FLOTAC®
35
The (?) enables the first four consecutive steps of the Mini-FLOTAC
technique, i.e. sample collection and weighing, homogenization, filtration and
filling.
Fill-FLOTAC®
36
The technique is a dilution technique in which 4 ml (~4 gm) of feces
(determined by displacement) is suspended in 56 ml of 0.1 N sodium
hydroxide.
D. STOLL'S EGG COUNTING TECHNIQUE
37
The sodium hydroxide saponifies fat and frees eggs from fecal debris.
D. STOLL'S EGG COUNTING TECHNIQUE
38
The amount of diluted stool used for egg counting is measured by Stoll
pipettes calibrated at 0.075 mL and 0.15 mL
D. STOLL'S EGG COUNTING TECHNIQUE
39
The constant used to multiply the total egg count depends on the amount of
stool examined.
D. STOLL'S EGG COUNTING TECHNIQUE
40
Materials and reagents:
Stool displacement flask etched lines at 56 and 60 ml.
Pipettes for 0.150 and 0.0.075 ml
0.1 N sodium hydroxide
Glass beads
Glass slide and coverslip
D. STOLL'S EGG COUNTING TECHNIQUE
41
1. In a calibrated Stoll flask, add 0.1 N sodium hydroxide to the (?).
56-ml mark
42
2. Add fresh fecal material to the flask so that the level of fluid rises to the 60-ml mark. This amount of feces is equivalent to (?).
4 g of feces
43
3. Add a few (?), and shake vigorously to make a uniform suspension. If the
specimen is hard, the mixture may be placed in a refrigerator overnight before shaking
to aid in mixing.
glass beads
44
4. With a calibrated pipette, quickly remove [?] (or 0.075 ml) of suspension and
transfer it to a slide.
0.15 ml
45
5. Do not use a (?); place the slide on a mechanical stage, and count all of the
eggs.
coverslip
46
6. Multiply the egg count by 100 if (1) stool suspension is used (or 200 if [2] is
used) to obtain the number of eggs per gram of stool.
1. 0.15 ml
2. 0.075 ml
47
7. The estimate (eggs per gram) obtained will vary according to the (?) of the stool.
48
The following correction factors should be used to convert the estimate to a formed-
stool basis:
mushy formed ........................... ?
mushy ......................................... ?
mushy diarrheic ......................... ?
1.5
2
3
49
Egg counts on liquid specimens are generally unreliable; the most accurate
counts are obtained with use of (??.
formed or semi-formed specimens
50
crude but useful semi-quantitiative method based on the assumption that an ideal fecal smear contains (?) of stool
1 to 2 mg
51
An estimated (?) stool in a standard smear is covered by 22 by 22 mm
coverslip and all eggs in the entire preparation are counted.
1 to 2 mg
52
The number of eggs per gram (EPG) is obtained by multiplying the total counts
by 667 if the standard smear has (?) stool (1000 mg divided by 1.5 )
1.5 mg
53
o the factor is 500 if the smear has (?) stool, etc. Obviously,
results are subject to large variation if the total egg count per
coverslip is low.
2 mg
54
1. Microscopic examination
2. Colonoscopy, Sigmoidoscopy, or Proctoscopy
3. Egg count test (Kato-Katz)
A. Trichuris trichiura
55
– detection of the characteristic eggs in stools
Microscopic examination
56
a. Direct fecal smear (DFS)
b. Kato thick smear (KTS) – not used in protozoans; ova
c. Kato-Katz
Trichuris trichiura Microscopic examination
57
o Modified kato-thick
Kato-Katz
58
o To count egg per gram
Kato-Katz
59
o With sheaving of stool (debris removal – more refined stool sample)
Kato-Katz
60
o To easily count parasite
Kato-Katz
61
1. Place paper
2. Place sheave on top of stool sample
3. Get a slide and place a template (board or plastic that gives
volume of stool being examined)
a. Thickness of 1,5mm
b. Diameter of hole: 6.5
c.
4. Fill out the hole on the slide when template is removed
Kato-Katz
62
o Stool concentation techniques (sedimentation and flotation) or Formalin-
ether acetate sedimentation
Kato-Katz
63
Adult worms
Colonoscopy, Sigmoidoscopy, or Proctoscopy
64
Adult worms – visible on macroscopic examination of the:
o intestinal mucosa.
o intestinal tract down to and including the rectum in heavy infections
65
identifying intensity of infection
o to determine the dose
o to follow-up effect of treatment
Egg count test (Kato-Katz)
66
1. Microscopic examination
2. Duodenal aspiration
Capillaria philippinensis
67
– recovery of the characteristic eggs in stools; larva; adult
Microscopic examination
68
Direct fecal smear (DFS)
Kato thick smear (KTS)
Kato-Katz
Stool concentation techniques (sedimentation and flotation) or Formalin-ether acetate
sedimentation
Capillaria philippinensis Microscopic examination
69
1. Histopathologic examination.
2. Serological test
Capillaria hepatica
70
Eggs - identified on the basis of their characteristic morphology on liver biopsy specimens
Histopathologic examination.
71
true human infection – no eggs are found in the stool
Capillaria hepatica
72
presence eggs in stool during (O&P) examinations – spurious infection following consumption of
egg-laden animal liver
Capillaria hepatica
73
unembryonated eggs – non-infectious
Capillaria hepatica
74
o merely pass through the digestive tract with feces
unembryonated eggs – non-infectious
75
IFAT
Serological test
76
– testing of human sera for the detection of early Capillaria hepatica infection
IFAT
77
1. Muscle biopsy
2. Beck’s Xenodiagnosis
3. Serological tests
4. Bachman intradermal test
5. Molecular methods
6. Radiological examination
7. Other laboratory findings
Trichinella spiralis
78
- demonstration of spiral larvae
Muscle biopsy
79
- Definitive diagnostic exam
Muscle biopsy
80
- can only be done when encystment of the parasite has occurred (7 days after infection)
Muscle biopsy
81
- Parasitized muscles
Muscle biopsy
82
diaphragm, pectoral gluteus, deltoid, biceps, & gastrocnemius
Parasitized muscles
83
At least 1 gram of muscle
Parasitized muscles
84
preferably near tendon insertion
Parasitized muscles
85
o chosen for taking diagnostic muscle biopsies
o easily accessible
diaphragm, pectoral gluteus, deltoid, biceps, & gastrocnemius
86
- Muscle fibers
Muscle biopsy
87
digested with trypsin
Muscle fibers
88
mounted on a glass slide
Muscle fibers
89
examined under microscope
Muscle fibers
90
o a teased preparation of muscle tissue is prepared in a drop of saline
solution
examined under microscope
91
o squeezed between two glass slides
examined under microscope
92
o muscle tissue is stained with safranin
examined under microscope
93
- diagnosis of trichinosis
Beck’s Xenodiagnosis
94
- Albino rats are fed with infected patient meat & are observed for around 14 days
Beck’s Xenodiagnosis
95
- rats are then killed in a month or so later and checked (particularly in the diaphragm) for the presence of
Trichinella spiralis larvae
Beck’s Xenodiagnosis
96
- Observe for the presence of female worms in the duodenum & the larvae in the muscles
Beck’s Xenodiagnosis
97
- rarely requested and is not available in most clinical laboratory
Beck’s Xenodiagnosis
98
- lab animal is examined
Beck’s Xenodiagnosis
99
- massive hypergammaglobulinemia with elevated serum IgE
Serological tests
100
- Confirmatory test
Serological tests
101
Bentonite flocculation test (BFT)
Latex flocculation test (LFT)
IFAT
ELISA
Trichinella spiralis Confirmatory test
102
– (+): recent infection
Bentonite flocculation test (BFT)
103
– - (+): recent infection
Latex flocculation test (LFT)
104
– T. spiralis antibody can be detected using TSL-1 secreting antigens obtained from the
infective stage larvae
ELISA
105
o Westernblot
o Latex agglutination
ELISA
106
- hypersensitivity test
Bachman intradermal test
107
- 1:5,000 or 1:10,000 dilution of the larval antigen
Bachman intradermal test
108
erythematous wheal (undulation) appears in positive cases within ?
15- 20 minutes
109
rest remains positive for (?) after infection
years
110
1. antigen is administered
rest remains positive for years after infection
111
PCR
Molecular methods
112
– species identification
PCR
113
X-ray
Radiological examination
114
– Calcified cysts may be demonstrated
X-ray
115
Eosinophilia and leukocytosis may also serve as indicators for disease.
Other laboratory findings
116
Elevated serum muscle enzyme levels, such as, may also aid in T. spiralis diagnosis.
Other laboratory findings
117
o lactate dehydrogenase
o aldolase
o creatine phosphokinase
o Myokinase w/ eosinophilia
Biochemical tests
118
– elevated enzymes as indicators for diagnosis
Biochemical tests
119
1. Microscopic identification of eggs
2. Macroscopic identification of adults
3. Serology
Ascaris lumbricoides
120
Definitive diagnosis of ascariasis
Microscopic identification of eggs
121
demonstration of the characteristic
fertilized and/or unfertilized eggs in stools
Microscopic identification of eggs
122
Reproductive rate: 200,000 eggs per day
Microscopic identification of eggs
123
Life span of adult worm: 1 year
Microscopic identification of eggs
124
a. Direct fecal smears (saline and
iodine-stained DFS)
b. Kato thick smear (KTS)
c. stool concentation techniques
(sedimentation and flotation)
d. Kato-Katz (KK) technique
Ascaris lumbricoides
Microscopic identification of eggs
125
Entangled mass in GIT cause obstruction
Macroscopic identification of adults
126
extraintestinal ascariasis
Ascaris lumbricoides Serology
127
a. Indirect hemagglutination (IHA)
b. Indirect fluorescent antibody
(IFA)
c. ELISA
Ascaris lumbricoides Serology
128
Sedimentation concentration test is
recommended instead of flotation.
Unfertilized eggs of Ascaris lumbricoides do not easily float.
Ascaris lumbricoides
129
Adult worms of Ascaris lumbricoides are occasionally passed in the stool or through the mouth or nose and are recognizable by
their macroscopic characteristics.
Ascaris lumbricoides
130
Intestinal disease can often be diagnosed
from radiographic studies of the gastrointestinal tract, in which the worm in
the intestinal tract may be visualized .
Ascaris lumbricoides
131
Larvae of Ascaris lumbricoides can be
identified in sputum or gastric aspirate
during the pulmonary migration phase
(examine fixed organisms for morphology).
Ascaris lumbricoides
132
1. Microscopic examination
Enterobius vermicularis
133
A. Graham’s scotch adhesive tape swab
B. NIH swab
Enterobius vermicularis Microscopic examination
134
- demonstration of eggs or
adult female worms
Graham’s scotch adhesive tape swab
135
- cellophane tape, or cellulose tape, or adhesive tape
Graham’s scotch adhesive tape swab
136
- Specimens
- perianal region
- early morning, before going to the toilet or bathing
Graham’s scotch adhesive tape swab
137
- Preferred
Graham’s scotch adhesive tape swab
138
- Yields the highest % of + results and the greatest no. of eggs seen
Graham’s scotch adhesive tape swab
139
- Placed on the perianal region
Graham’s scotch adhesive tape swab
140
MATERIALS
Clear cellulose tape
Microscope slide
Tongue depressor or a similar material
Graham’s scotch adhesive tape swab
141
1. Place a [?] (adhesive side
down) on a microscope slide as follows:
strip of clear cellulose tape
142
starting at (?) from one end,
run the tape toward the same end
1.5 cm
143
wrap the tape (?) to
the opposite end
around the slide
144
? even with the end of
the slide.
Tear the tape
145
? to the tape at the
end torn flush with the slide.
Attach a label
146
2. To obtain a sample from the perianal area, (?) by gripping the labeled end, and, with the
tape looped (adhesive side outward) over a wooden
tongue depressor that is held firmly against the slide
and extended about 2.5 cm beyond it, press the tape
firmly several times against the right and left perianal
folds.
peel back
the tape
147
3. Smooth the tape back on the slide, (?) down.
adhesive side
148
4. Label with ?.
patient name and date
149
5. Submit the (?) to the laboratory in a plastic bag.
tapes and slides
150
6. Examine the slide directly under the (?) of the microscope. To make the eggs more
visible, lift the tape from the slide and add a drop of
xylene or toluene to the slide. Press the tape back
down on the slide. Examine with low light intensity
on low power.
low-power objective
151
- named after National Institutes of Health, United States of America
(USA
NIH swab
152
consists of a glass rod at one end of which a piece of
transparent cellophane is attached with a rubber band
NIH swab
153
glass rod is fixed on a rubber stopper and kept in a wide test
tube
NIH swab
154
cellophane part is used for swabbing by rolling over the
perianal area
NIH swab
155
It is returned to the test tube and sent to the laboratory,
where the cellophane piece is detached, spread over a glass
slide and examined microscopically
NIH swab
156
Eggs are rarely found in the stool (in only about 5% of
infected persons) because release is external to the
intestine. Therefore, stool is NOT the specimen of
choice.
Enterobius vermicularis
157
The perianal swabs may also be collected late in the
evening, when the patient has been sleeping for several
hours.
Enterobius vermicularis
158
The eggs may sometimes be demonstrated in the dirt
collected from beneath the finger nails in infected
children.
Enterobius vermicularis
159
Care must be taken during specimen collection and
examination because eggs, if present, are already
infectious.
Enterobius vermicularis
160
A minimum of four to six consecutive negative slides is
required to rule out the infection.
Enterobius vermicularis
161
Alternatively, commercial “Swube tubes” (a paddle
coated with adhesive material) can also be used for
collection.
Enterobius vermicularis
162
1. Microscopic examination
2. Coproculture
Hookworms
163
– demonstration of the characteristic hookworm
eggs in stools
Hookworms Microscopic examination
164
a. direct fecal smears (saline and iodine-stained DFS)
b. Kato thick smear (KTS)
c. stool concentation techniques (sedimentation and flotation)
d. Kato-Katz (KK)
Hookworms Microscopic examination
165
Eggs of N. americanus and A. duodenale cannot be differentiated by morphology, thus, is reported as" hookworm species eggs"
Hookworms
166
If the stool remains unpreserved for over 24 h, the eggs may continue
to develop and the larvae may hatch. These larvae must be
differentiated from those of Strongyloides stercoralis (will be
discussed later), since therapies for the two infections are different.
Hookworms
167
Kato-Katz or kato thick smear preparation must be examined within 1 hour after preparation. Allowing the slides to stand for a long period
of time will cause drying and thin shells of hookworm egg will become
too transparent and will be difficult to see.
Hookworms
168
Adult hookworms are rarely seen in feces but may be recovered
following treatment. Examination of the buccal capsule is necessary to
determine the specific hookworm species.
Hookworms
169
Harada-Mori technique
Coproculture
170
- use of certain fecal culture methods (sometimes referred to as coproculture) is
especially helpful for detection of light infections with hookworm (also Strongyloides
stercoralis and Trichostrongylus spp)
Coproculture
171
- is also carried out to demonstrate larvae from feces which helps in distinguishing N.
americanus and A. duodenale
Coproculture
172
- filter paper test tube culture technique
Harada-Mori technique
173
- initially introduced by Harada and Mori in 1955
and was later modified by others
Harada-Mori technique
174
- The technique employs a filter paper to which fecal material is
added and a test tube into which the filter paper is inserted
Harada-Mori technique
175
- Moisture is provided by adding water or saline to the tube.
Harada-Mori technique
176
- The water continuously soaks the filter paper by capillary action.
Harada-Mori technique
177
- Incubation under suitable conditions favors hatching of ova
and/or development of larvae.
Harada-Mori technique
178
Appropriate PPE (gloves, gown) should always be worn to minimize
the risk of transdermal penetration when working with stool
specimens or larval cultures.
Hookworms
179
Fecal specimens to be cultured should not be refrigerated, since some
parasites (especially Necator americanus) are susceptible to cold and
may fail to develop after refrigeration.
Hookworms
180
Rhabditiform (L1) larvae that hatch from eggs are 250-300 µm long
and approximately 15-20 µm wide. They have a long buccal canal and
an inconspicuous genital primordium. Rhabditiform larvae are usually
not found in stool, but may be found if there is a delay in processing
the stool specimen. If larvae are seen in stool, they must be
differentiated from the L1 larvae of Strongyloides stercoralis.
Hookworms
181
Filariform (L3) larvae are 500—700 µm long. They have a pointed tail
and are ensheathed, with esophagus that is one-third the length of
the body. Some subtle morphological differences exist between N.
americanus and A. duodenale at this stage. These L3 are found in the
environment and infect the human host by penetration of the skin..
Hookworms
182
Diagnosis is based mainly on history and clinical manifestations.
Zoonotic hookworms
183
1. Microscopic examination
2. Coproculture
3. Serology
4. Molecular diagnosis
Strongyloides stercoralis
184
a. Direct stool microscopy
b. Baermann technique
c. Microscopic examination of duodenal and respiratory specimens
Strongyloides stercoralis Microscopic examination
185
Demonstration of the rhabditiform larvae in freshly passed stools
Direct stool microscopy
186
most important method of specific diagnosis
Direct stool microscopy
187
may be recovered in direct fecal smears or stool concentration techniques
Direct stool microscopy
188
It may be difficult to observe morphologic details in rapidly moving larvae; a drop of iodine or formalin or slight heating can be used to
kill the larvae.
Strongyloides stercoralis
189
Larvae found in old, unpreserved stools have to be differentiated from larvae hatched from hookworm eggs.
Strongyloides stercoralis
190
In addition to rhabditiform larvae, eggs of S. stercoralis, often indistinguishable from those of the hookworm, may be present in stool
samples from patients suffering from severe diarrhea.
Strongyloides stercoralis
191
Note that formalin-ethyl acetate concentration may remove larvae and reduce sensitivity. Sedimentation methods have a typically
higher recovery rate than those in which a flotation technique has been used.
Strongyloides stercoralis
192
Baermann technique is another sedimentation method of examining a stool specimen suspected of containing small numbers of
Strongyloides larvae.
Baermann technique
193
This uses a funnel apparatus and relies on the principle that active larvae will migrate from a fresh fecal specimen that has been
placed on a wire mesh with several layers of gauze which are in contact with tap water.
Baermann technique
194
Larvae migrate through the gauze into the water and settle to the bottom of the funnel, where they can be collected and examined.
Baermann technique
195
Specimens that have been refrigerated or preserved are not suitable for culture. Larvae of certain species are susceptible to cold
environments.
Strongyloides stercoralis
196
Besides being useful for diagnosis from stool specimens directly, it can be enhanced by culture if no larvae are seen in 2-3 hours.
Strongyloides stercoralis
197
Infective larvae may be found any time after the fourth day and occasionally after the first day in heavy infections. Caution must be
exercised in handling the fluid, gauze pad, and beaker to prevent infection. Wear gloves when using this technique.
Strongyloides stercoralis
198
Make sure that the end of the tubing is well inside the beaker before releasing the pinch clamp. Release the pinch clamp slowly to
prevent splashing.
Strongyloides stercoralis
199
Besides being used for patient fecal specimens, this technique can be used to examine soil specimens for the presence of larvae.
Strongyloides stercoralis
200
Beale’s string test
Microscopic examination of duodenal and respiratory specimens
201
Duodenal aspiration
Microscopic examination of duodenal and respiratory specimens
202
Aspiration of duodenal fluid or use of the less invasive Entero-test (commonly called a string test) may be useful to detect larvae in
patients with negative stool samples.
Microscopic examination of duodenal and respiratory specimens
203
In disseminated strongyloidiasis, filariform larvae may be detected in sputum, bronchial washings or pleural fluid.
Microscopic examination of duodenal and respiratory specimens
204
Principle:
o Active larvae will migrate out of a fecal specimen that has been placed on a wire mesh covered with several layers
of gauze.
Microscopic examination of duodenal and respiratory specimens
205
a. Harada-Mori technique
b. Agar plate cultures
c. Charcoal culture method
Strongyloides stercoralis
Coproculture
206
- discussed previously in hookworm species.
Harada-Mori technique
207
These are also recommended for the recovery of S. stercoralis larvae and tend to be a more sensitive method than the usual direct
smear or fecal concentration methods.
Agar plate cultures
208
Stool is placed onto agar plates, and the plates are sealed to prevent accidental infections and held for 2 days at room temperature.
Agar plate cultures
209
As the larvae crawl over the agar, they carry bacteria with them, thus creating visible tracks over the agar .
Agar plate cultures
210
The plates are examined under the microscope for confirmation of the presence of larvae.
Agar plate cultures
211
Another way to culture hookworm, Strongyloides, and trichostrongyle larvae is by using a granulated charcoal culture.
Charcoal culture method
212
The conditions of this culture provide an environment for larval development that mimics conditions in nature.
Charcoal culture method
213
It provides an efficient way to harvest large numbers of infective-stage larvae for use in experimental infections.
Charcoal culture method
214
Most antibody detection tests employ antigens derived from Strongyloides stercoralis (or from closely-related S. ratti or S.
venezuelensis) filariform larvae.
Strongyloides stercoralis Serology
215
Although indirect fluorescent antibody (IFA), indirect hemagglutination (IHA) and antigen-linked fluorescent and magnetic bead tests
are are available, enzyme immunoassay (EIA) is recommended because of its greater sensitivity.
Strongyloides stercoralis Serology
216
The commercial EIA kits that are currently available have sensitivity of 95%.
Strongyloides stercoralis Serology
217
PCR and LAMP methods may be used to detect strongyloidiasis in fresh, frozen, or non-formalin fixed stool specimens.
Strongyloides stercoralis
Molecular diagnosis
218
Sensitivity and specificity vary depending on the reference test used to calculate such characteristics; false negatives and false
positives do occur.
Strongyloides stercoralis
Molecular diagnosis
219
Diagnosis of toxocariasis relies mostly on indirect means,
particularly serology, since larvae are trapped in tissues and
not readily detected morphologically.
Toxocara spp.
220
While visualization of larvae in histologic sections provides
unequivocal diagnosis, the probability of capturing a larva in a
small biopsy specimen is low.
Toxocara spp.
221
For both VLM and OLM, a presumptive diagnosis rests on
clinical signs, compatible exposure history, laboratory
findings (including eosinophilia), and the detection of
antibodies to Toxocara.
Toxocara spp.
222
Detection of microfilariae in blood is the traditional method of diagnosis of most filarial nematode infections, which include those caused by W. bancrofti, Brugia species, L. loa, M. perstans, and M. ozzardi.
FILARIA
Microscopy
223
The notable exception to the use of blood specimens is for the diagnosis of onchocerciasis and M. streptocerca infection, which involves examination of skin snips.
FILARIA
Microscopy
224
For more sensitive detection of microfilariae in blood, it is important to understand the periodicity of a given species or geographical variant.
Blood examination
o Periodicity
225
This determines the optimal time their microfilariae will be circulating in peripheral blood and, therefore, the best time of the day to obtain blood specimens for examination.
Blood examination
o Periodicity
226
To determine microfilarial periodicity in an individual, it is necessary to examine measured quantities of blood collected at consecutive intervals of 2 or 4 hours over a period of 24-30 hours.
Blood examination
o Periodicity
227
Methods used for the detection of microfilariae in the blood
i. Stained blood films
ii. Wet mount of capillary blood
iii. Microhematocrit method
iv. Quantitative buffy coat (QBC) method
v. Concentration procedures
228
i. Stained blood films
Preparation of thick blood films
Staining of thick blood films
Microscopic examination
229
The examination of thick blood films is the most widely used method in field surveys of filarial infection.
Preparation of thick blood films
230
Carefully measured blood samples of at least 20 µl and preferably 60 µl in volume are recommended.
Preparation of thick blood films
231
At least two thick films should be made. Identification of microfilariae in thin films is not recommended because the microfilariae are shrunken, distorted and difficult to recognize.
Preparation of thick blood films
232
Giemsa and Delafield's hematoxylin are the preferred and most widely used for preparing permanently-stained thick blood films in the detection of microfilariae.
Staining of thick blood films
233
With Giemsa stain, nuclei of microfilariae will stain blue to purple, and the sheath will stain pink (B. malayi).
Staining of thick blood films
234
Hematoxylin enhances nuclear detail in microfilaria, and stains the sheath greyish-blue (W. bancrofti, B. timori, L. loa).
Staining of thick blood films
235
Because microfilariae are large, they can be detected by screening at a lower magnification.
Microscopic examination
236
The entire thick blood film should be examined with the 10x low-power objective.
Microscopic examination
237
When microfilariae are seen, morphologic details can be seen using the 40x high dry objective, and then examined at a higher magnification (×100 with oil) for species-level identification.
Microscopic examination
238
A drop of blood directly on a slide is added with an equal-sized drop of NSS, and examined under the microscope (10x objective).
ii. Wet mount of capillary blood
239
Microscopic examination of fresh blood has limited usefulness.
ii. Wet mount of capillary blood
240
The whiplike motion of the microfilariae are easily spotted because they cause movement of surrounding red blood cells, but species identification is not possible.
ii. Wet mount of capillary blood
241
However, in regions where only one species is found, its presence and density in the blood can be determined with reasonable accuracy by this means.
ii. Wet mount of capillary blood
242
PROCEDURE:
(1) Three-quarters fill a microhaematocrit capillary tube with the blood. Seal one end of the tube.
(2) Centrifuge in a microhaematocrit centrifuge at 10 000g for 2 minutes.
(3) Lay the capillary tube on a slide and secure the two ends with adhesive tape.
(4) Examine the dividing line between the blood cells and the plasma under the microscope using the 10x objective.
Motile microfilariae will be seen at the bottom of the column of plasma, just above the layer of leukocytes and erythrocytes.
The tube can be snapped at the bottom of the column of plasma.
Use the first drop from each piece of the broken tube to prepare a thick film.
Stain the film as to identify the species.
iii. Microhematocrit method
243
(1) ? fill a microhaematocrit capillary tube with the blood. Seal one end of the tube.
Three-quarters
244
(2) Centrifuge in a microhaematocrit centrifuge at ?.
10 000g for 2 minutes
245
(3) Lay the capillary tube on a slide and secure the two ends with ?.
adhesive tape
246
(4) Examine the (?) between the blood cells and the plasma under the microscope using the 10x objective.
dividing line
247
? will be seen at the bottom of the column of plasma, just above the layer of leukocytes and erythrocytes.
Motile microfilariae
248
The tube can be snapped at the bottom of the ?.
column of plasma
249
Use the (?) from each piece of the broken tube to prepare a thick film.
first drop
250
? as to identify the species.
Stain the film
251
The utilization of quantitative buffy coat tube coated with acridine orange (discussed in malaria diagnosis) has been reported to be an acceptable rapid diagnostic test for detection of microfilariae, with a sensitivity equivalent to that of thick blood film.
iv. Quantitative buffy coat (QBC) method
252
v. Concentration procedures
Knott's concentration method
Membrane filtration technique
253
These procedures are preferred when the microfilarial load is low.
v. Concentration procedures
254
They can be used for the enhanced detection of microfilariae in blood or other body fluids.
v. Concentration procedures
255
PROCEDURE:
(1) Collect 1 ml of blood (whole or citrated) by venipuncture and place in a 15-ml centrifuge tube containing at least 10 ml of 2% formalin; stopper the tabe and shake vigorously. The formalin lyses RBCs, fixes blood protozoa, and kills and straightens the bodies of microfilariae.
(2) Centrifuge at at approximately 300 g for 2 minutes. If a centrifuge is not available, place the tube in an upright position for 12 hours for gravitational sedimentation.
(3) Decant the supernatant fluid (the small amount remaining in the tube is allowed to flow back on to the sediment).
(4) Remove the sediment with a Pasteur pipette and examine a drop of sediment on a slide under a coverslip with the low power objective of the microscope.
(5) A portion of the sediment may be spread on a glass slide as a thick film and allowed to dry thoroughly. Stain the film with Giemsa or hematoxylin stain.
Knott's concentration method
256
(1) Collect (?) of blood (whole or citrated) by venipuncture and place in a 15-ml centrifuge tube containing at least 10 ml of 2% formalin; stopper the tabe and shake vigorously. The formalin lyses RBCs, fixes blood protozoa, and kills and straightens the bodies of microfilariae.
1 ml
257
(2) Centrifuge at at approximately (?). If a centrifuge is not available, place the tube in an upright position for 12 hours for gravitational sedimentation.
300 g for 2 minutes
258
(3) Decant the [?] (the small amount remaining in the tube is allowed to flow back on to the sediment).
supernatant fluid
259
(4) Remove the sediment with a Pasteur pipette and examine a drop of sediment on a slide under a coverslip with the (?) of the microscope.
low power objective
260
(5) A (?) may be spread on a glass slide as a thick film and allowed to dry thoroughly. Stain the film with Giemsa or hematoxylin stain.
portion of the sediment
261
This allows for removal of elements by filtration through a membrane of desired pore size.
Membrane filtration technique
262
Cellulose-mixed-ester filters (e.g., Millipore filters) and polycarbonate filters (e.g., Nucleopore filters), 25 mm in diameter, are most common filters used.
Membrane filtration technique
263
Although a membrane filter of 5-µm porosity is ideal for isolation of most microfilariae in the blood, it is unsatisfactory for the isolation of Mansonella microfilariae because of their small size. A 3-µm-pore-size filter could be used for recovery of this organism.
Membrane filtration technique
264
PROCEDURE:
(1) Collect fresh blood in sodium citrate or EDTA
(2) Add 1 ml of the blood to 10 ml of 10% Teepol-saline solution (Teepol lyses the red blood cells;
prepared by adding 50 g Teepol concentrate to 450 ml saline).
(3) Place a membrane filter over supporting filter paper moistened with water securely into a filter
holder.
(4) Remove plunger from the barrel of a 20-ml syringe and connect the barrel of the syringe to the
filter holder.
(5) Pour the blood-Teepol mixture (see step 2) into the barrel of syringe, replace the plunger in the
syringe, and gently force the solution through the filter.
(6) Remove the syringe from the filter holder, draw up 10 ml of water into syringe, reattach the
filter holder, and gently wash the filter by flushing the water through it.
(7) Force two syringe-volumes of air through filter to expel excess water and make microfilariae
more adherent to filter.
Membrane filtration technique
265
(1) Collect fresh blood in sodium citrate or ?
EDTA
266
(2) Add 1 ml of the blood to (?) Teepol-saline solution (Teepol lyses the red blood cells;
prepared by adding 50 g Teepol concentrate to 450 ml saline).
10 ml of 10%
267
(3) Place a (?) over supporting filter paper moistened with water securely into a filter
holder.
membrane filter
268
(4) Remove plunger from the barrel of a (?) and connect the barrel of the syringe to the
filter holder.
20-ml syringe
269
(5) Pour the (?) (see step 2) into the barrel of syringe, replace the plunger in the
syringe, and gently force the solution through the filter.
blood-Teepol mixture
270
(6) Remove the syringe from the filter holder, draw up (?) into syringe, reattach the
filter holder, and gently wash the filter by flushing the water through it.
10 ml of water
271
(7) Force (?) of air through filter to expel excess water and make microfilariae
more adherent to filter.
two syringe-volumes
272
(8) Pass (?) through the filter to fix the microfilariae.
3 ml of methanol
273
(9) Remove the (?) from the holder and place on a glass slide; allow to dry thoroughly.
filter
274
(10) Stain with (?) per the normal procedure.
Giemsa or hematoxylin stain
275
(11) Dip the slide in (?) to avoid air bubble formation.
toluene
276
(12) Add a drop of (?) and a coverslip.
mounting medium
277
(13) Examine the slide when ?.
dry
278
? is the gold standard test used for the detection of microfilariae of Onchocerca volvulus and Mansonella streptocerca.
Skin snip examination
279
A very small piece of the patient’s skin is collected. The skin snips are generally collected from several sites on the body to maximize detection sensitivity. The preferred site is the skin over bony prominences of the iliac crest (top of the buttocks) and the scapular area (shoulder blades), and in lower calves (legs). If the patient has nodules, the specimen is taken from the skin i n t h e c e n t e r o f t h e n o d u l e . I t i s recommended that six
specimens (two snips from all three of these sites on each side of the body of the individual) should be examined before reporting a negative result.
Skin snip examination
280
The specimen should not be bloodstained. When snips are cut too deeply, small capillaries may be lacerated and the snip may be contaminated by microfilariae that might be present in the patient's blood.
Skin snip examination
281
Skin snips are obtained in one of two ways:
1. Using a sterile needle
2. Using a sclerocorneal biopsy punch
282
Pierce the skin to a depth of 2-3 mm, and elevate a piece of skin with the point of the needle and shave it off with a scalpel, as close to the needle as possible.
Using a sterile needle
283
This takes snips of uniform diameter (approximately 2.3-2.5 mm)
Using a sclerocorneal biopsy punch
284
The skin snips are examined under the microscope as:
1. Wet mount preparation
2. Permanent stained preparation
285
o Place the fragment of skin in the drop of NSS on the slide. Do not flatten the piece of skin; if only one microfilaria is present, it might be damaged.
o Cover with a coverslip. If any part of the specimen is not in the coverslip with a Pasteur pipette, until the whole area underneath the coverslip is wet.
o After 30 minutes to 3 hours, examine t h e p r e p a r a t i o n u n d e r t h e microscope using the 10x low power objective.
Microfilariae are highly motile. If any are present, they will be seen moving towards the NSS.
If negative, continue incubating overnight at 37°C.
Wet mount preparation
