This document provides information on various special stains used to identify microorganisms and cellular structures under the microscope. It discusses stains used for flagella, metachromatic granules, spirochetes, Chlamydia, rickettsia, and fungi. Specific stains covered include Wright's Giemsa, Gram, acid-fast, silver, toluidine blue, calcofluor white, acridine orange, auramine phenol, and lactophenol cotton blue. Procedures for each stain are provided along with what structures or organisms they help identify.

1. DR.YOGITA Mistry
GMC,Surat

2. Name of special stains
 Acridine orange
 Calcofluor white
 Aura mine phenol
 Toluidine blue
 Wright’s – Giemsa stain
 Albert’s stain
 LPC

3.  Stains for flagella
 Leif son stain
 Gray method
 Silver technique
 Stains for metachromatic granules
 Albert stain
 Neisser stain
 Leoffler methylene blue
 Ponder’s stain
 Stains for spirochetes
 Fontana
 Levaditi
 Silver Impregnation
 India ink negative stain

4.  For Chlamydia
 Gimenez
 Machiavello
 Giemsa
 For rickettsia
 Castaneda
 Machiavello
 Giemsa

5. WRIGHT’S GIEMSA STAIN
 Mainly stains the cellular eliments of the peripheral
blood , has little use for staining bacteria.
 Used primarily to detect the intracellular yeast form of
Histoplasma capsulatum or intracellular amastigotes
of Leishmania species or Trypanosoma cruzi, malarial
parasites.
 Also helpful for demonstrating certain intracellular
viral inclusions.
 There are two ways of staining-rapid and slow method.

6.  RAPID METHOD:
 Useful for staining the malarial parasite, schuffener’s
dots and also for trypanosoma demonstration.
 Method:
 Fix films in methyl alcohol for 3 min.
 Stain in a mixture of 1 part stain and 10 parts buffer
solution for 1 hr.
 Wash with buffer solution(buffered water ,ph 7-7.2),
allowing preparation to differentiate for 30 seconds.
 Blot and allow to dry in air.

7.  Rapid method with heat application is useful for
demonstrating spirochetes.
 Method:
 Fix the smear with absolute alcohol foe 15 min. or by
drawing three times through flame.
 Cover a fixed film with the diluted stain. warm till
steam rises , allow to cool for 15 seconds, then pour off
and replace with fresh stain and heat again.
 Repeat the procedure 4-5 times , wash in distilled
water , dry and mount.

8. Histoplasm
cysts

10. Ring form of P . falciparum

11.  SLOW METHOD:
 Useful for demonstrating objects difficult to stain in
the ordinary way such as spirochetes.
 Principle is to allow the diluted stain to act for a
considerable period.
 But care has to be taken as fine precipitate can de
deposited.
 Procedure:
 Fix the film in methyl alcohol for 3 min.
 Mix 1 ml stain with 20 ml buffer solution.
 Place a piece of thin glass rod in the stain in the dish.
 Leave to stain for 16-24 hr.
 Wash the slide for in a steam of buffer solution.
 Allow to dry in air , mount and examine.

12. spirochetes

13. STAINS FOR METACHROMATIC GRANULES
 Albert stain
 Neisser stain
 Ponder’s stain
 Loeffler’s methyline blue
 ALBERT’S STAIN
Albert’s staining is specially demonstrates the presence or
absence of the metachromatic granules, a characteristic
feature of Corny bacterium diphtheria. During gram staining
if a smear appears as purple rods with straight or slightly
curved with clubs at the end, with a characteristic V shape
then it is suspected as Corny bacterium diphtheria. The
further confirmation can be done by Albert’s staining
technique

14. ALBERT’S STAIN
 . This techniques employ two stains:
 Albert’s solution
 Toluidine blue 1.5 gm
 Malachite green 2gm
 Glacial acetic acid 10ml
 Alcohol 20ml
 Distilled water 1 liter

15.  Albert’s iodine
 Iodine 6gm
 Potassium iodide 9gm
 Distilled water 900ml
 Procedure
 Cover a heat fixed slide with Albert's stain and allow to act
for 3-5 min.
 Wash in water and blot dry.
 Cover with Albert's iodine for 1 min.
 Wash and dry.
 Granules stain bluish black , protoplasm green and other
organisms light green.

17.  Neisser's Stain
 Method
 Stain with Neisser's methylene blue for 3 min.
Rinse rapidly with water.
Stain with dilute iodine for 1 min.
Wash rapidly in water.
 Counter stain with neutral red for 1 min.
 Results: Cytoplasm appears pink and granules deep
blue

18.  Reagents
 Neisser’s stain solution
For use, mix 20 ml of solution A and 10 ml of solution B
 Neisser’s Solution A:
Methylene blue 0.1 gm;
95% ethanol 5 ml;
glacial acetic acid 5 ml
distilled water 100 ml.
 Dissolve the dye in the water and add the acid and ethanol
 Neisser’s Solution B:
Crystal violet 0.33 gm;
95% ethanol 3.3 ml
distilled water 100 ml.
 Dissolve the dye in the ethanol- water mixture

20. LPC STAIN FOR FUNGUS
 Fungi can be stained with lacto phenol cotton blue.
 Composition:
 The composition of lacto phenol cotton blue is as follows:
 Phenol crystals 20g
 Lactic acid 20 ml
 Glycerol 40 ml
 Distilled water 20 ml
 Cotton blue (or methyl blue) 0.075 g
 A drop of 95% alcohol is applied on the "slide. The fragments of the
culture of the fungus is placed on the slide. It is teased out gently with
needles. When it has spread well on the slide, allow the alcohol to
evaporate. Then add a drop of the stain ( lacto phenol cotton blue).
 A cover slip is applied carefully without allowing air bubbles to form. A
gentle pressure is applied and the excess stain around the cover slip is
removed with the edge of a blotting paper. Now the slide can be viewed
under the microscope to see the details of the fungus.

22. TOLUIDINE BLUE
 Mainly use for identification of cysts of Pneumocystis
jiroveci ( carinii).
 Reagents:
 Sulfation reagent;
 45 ml of glacial acetic acid is poured into a Coplin jar
which had been placed into a plastic tub filled with cool tap
water (not below 10°C).
 A 15-ml portion of concentrated sulfuric acid is slowly
added with a glass pipette, being careful not to produce
splashing.
 The solution is gently mixed with a glass rod. The sulfation
reagent can be kept at room temperature and could be used
for 1 week. .

23.  Toluidine blue 0 solution:
 0.3 g of Toluidine o
 60 ml of distilled water
 2.0 ml of concentrated hydrochloric acid
 140 ml of absolute ethyl alcohol.
 The solution is stored at room temperature and can be
used for 1 year. Staining was performed with all
solutions in Coplin jars.
 For other reagents:
 two Coplin jars each is needed for 95% ethyl alcohol,
absolute ethyl alcohol, xylene.

24.  (i) With forceps, slides are placed in the sulfation reagent for 10min. The
reagent is mixed with a glass stirring rod immediately after insertion of the
slides and again after 5 min.
 (ii)The slides removed from the sulfation reagent with forceps, placed in a
glass slide holder, and washed gently under cold running tap water for 5
min. The excess water is then drained.
 (iii) The slides are placed in toluidine blue 0 for 3 min.
 (iv) The slides are dipped in and out of 95% ethyl alcohol (in two Coplin
jars) for approximately 10 mins until clean, with most of the blue dye being
removed in the
 first jar.
 (v) The slides were dipped in and out of absolute ethyl alcohol (in two
Coplin jars) for approximately 10 mins for further decolorizing.
 (vi) The slides are dipped in and out of xylene (in two Coplin jars) for
approximately 10mins until clean.
 (vii) The back and frosted areas of the slides were wiped dry with a paper
towel.
 (viii)The slides are examined with 20x and 40x objectives; examination
under oil was is not necessary but could be done once the cover-slipped
slides had dried. The staining procedure itself took approximately 20 min.
Up to 4 slides could be stained readily simultaneously.

27. Calcofluor white stain
 Colorless dye
 Valuable flurochrome stain for rapid detection of fungi
in wet mounts, smears , and tissue sections.
 it fluorescence when exposed to long wave length UV
and short wave length visible light.
 Most use full in detecting yeast cell , hyphae ,
pseudohyphae in skin and mucous membrane
scraping
 Fungal structure will display a brilliant apple green or a
blue white ,depending on the wave length of the
exciter light and filter combination

28. Preparation of stain:
 Stain is prepared by dissolving 1 gm of flurochome in
100 distilled water.
 From this a working solution is made by diluting 10 ml
in 90 ml of 0.05% Evan's blue stain.
 For use, 1 drop is mixed with 1 drop of 20% KOH.

30. ACRIDINE ORANGE STAIN
 This stain is now use with increase frequency to detect
bacteria in smears prepare from fluids and exudates in
which bacteria are expected to be in low concentration.
 Use:
 It has been recommended for rapid identification of
trichomonas vaganalis ,yeast cells , clue cells in vaginal
smears.
 Also helpful to detect intra cellular gonococci ,
meningococci , and bacteria perticularly in blood cultures.
 Use of AO for staining acid fast bacilli and examined under
UV light can be more rapid and efficient screening method.
With UV light, RNA component orange red and DNA
component yellow-green seen.

31.  To make 500 ml of acridine orange acid stain:
 Acridine orange 0.13gm
 Acetic acid , glacial acid 10 gm
 Distilled water 490 ml
 Alcohol saline solution:
 Ethanol/methanol 5ml
 Physiological saline 245ml
 Method:
 Cover the unfixed dried smear with acridine orange acid stain for 5-10
seconds. This stain also fix the slide.
 Wash off the stain and decolorize the smear with alcohol saline
solution for 5-10 seconds.
 Rinse the smear with physiological saline and place the slide in
draining rack.
 Add a drop of saline or distilled water to the smear and cover with a
cover glass.
 Examine by using florescence microscope with BG 12 exciter filter and
No.44 and No.53 barrier filters. Alternatively ,use LED fluorescence.

32.  T. vaganalis orange red with yellow green nucleus
 Yeast cell orange
 Bacteria orange
 Leucocytes yellow green
 Epithelial cells yellow green

33. AURAMINE PHENOLE STAIN
 Mainly useful for detection of M.TB in sputum.
 As the smear can be examine at lower magnification ,
shorter time is required for a slide than Z N technique.
 Also few AFB are more likely to get detected by this
stain.
 With recent development of LED florescence system ,
I is now possible to examine smear inexpensively and
easily in district labs.
 When required, aura mine stained smears can be
restained by Z N stain by first treating the smear with
5% oxalic acid for 2 mins followed by washing in water.

34.  Aura mine phenol stain :
 Phenol crystals 15gm
 Aura mine o 1.5gm
 Distilled water 500 ml
 1%acid alcohol
 Ethanol/methanol 693ml
 Distilled water 297ml
 HCL 10ml
 Potassium permanganate 0.5 gm in 500 ml distilled
water t0 make 0.1% of it.

35.  Cover heat fixed dried smear with the filtered aura
mine phenol stain for 15 min.(this stain binds to
mycolic acid of TB bacilli)
 Wash with clean water.
 Decolorize with acid alcohol for 3-5 minutes.
 Wash off with clean water.
 Cover the smear with potassium permanganate
solution for 15 seconds and clean with water.(provides
the dark back ground)
 Tubercle bacilli fluoresce bright-yellow against a dark
back ground.

37.  Reporting of aura mine stained sputum smear
examined using 10 and 40 * objectives based on WHO
system:
 1-19 bacilli in 40 fields scanty
 20-199 bacilli in 40 fields +
 5-50 bacilli per field ++
 More than 50 bacilli per field +++

38. LOEFFLER METHYLENE BLUE
STAIN
Use full stain for identification of gram negative organism
such as H. influenza and N.meningitidis which often do
not stand out against red staining background of gram
stain.
 It stains the polymorph nuclear cells blue and bacterial
cells deep blue , back ground light gray in color.
 This stain should be consider as an adjunct to gram’s stain
in laboratories where inaccessibility to a fluroscence
microscope present.
 It is also helpful in respiratory secretion to detect
Pneumocystis jiroveci( carinii ) rapidly.
 Helpful to see PMN cells in inflammatory diarrheal disease
and mononuclear cells in stool in pt of typhoid.

39.  Methylene blue 0.5 gm
 Ehtanol 30ml
 Potassium hydroxide 0.1ml
 Distilled water 100ml
 Procedure:
 Place a drop of methylene blue stain on a slide. Mix a
small amount of specimen with the stain , cover with a
cover slip.
 Examine under 40* objective for leucocytes , pus cell,
RBCS.

40. NEW METHYLENE BLUE
 New methylene blue 1gm
 Sodium citrate 0.6gm
 Sodium chloride 0.7gm
 Distilled water 100ml
 It will detect the presence or absence of bacteria , their
number and shape.

41. Iodine preparation for stool
sample
 The value of wet preparations lies in the fact that certain
protozoa trophozoites retain their motility which may aid
in their identification. Definitive identification however
may not be possible, especially for amoeba, since the nuclei
of trophozoites and cysts are often not clearly visible. Wet
preparations on fresh unpreserved liquid stool should be
performed and examined as soon as possible (within 30
minutes of passage)
 Reagents
 Normal saline (0.85%)
Lugols iodine
 potassium iodide 10gm
 powdered iodine crystals 5gm
 distilled water 100ml

43. Eosin stain:
 Eosin powder 0.5 gm
 Distilled water 100ml
 Procedure:
 Place a drop of fresh physiological saline on one end of a slide
and a drop of eosin stain on other side.
 Using a piece of stick or wire loop , mix a small amount of fresh
specimen with each drop.
 Cover each preparation with cover slip.
 Smear should not be much thick otherwise amoebae or cysts
will not be seen.
 Examine under 10 and 40* objectives.
 Useful to see motile E.histolytica trophozoites containg red cells
, motile G. lamblia trophozoites , stongoloides larvae, eggs and
cysts of parasite.

45. FLAGELLAR STAIN
 Because of their extreme thinness, flagella are best
demonstrated with the electron microscpoe,metal
shadowed films or film made with phosphotungstic acid
for negative staining.
 But when electron microscope are not available , it is
possible to demonstrate the presence and arrangement of
flagella by special staining methods for the light
microscope.
 To be resolvable, flagella must be thickened at least 10 fold
by a superficial deposition of stain procured by the action
of mordant, usually tannic acid.
 An easier , cleaner and more reliable method is wet mount
flagellar stain.

46.  WET MOUNT FLAGELLAR STAIN:
Mordant solution:
 5% phenol 10 ml
 Tannic acid 2gm
 Aluminum potassium sulphate, 10 ml
Saturated aqueous solution
Ryu’s stain
Crystal violet , saturated ethanoic solution 10ml
Mordant solution 100ml

47. Procedure:
 After growing bacteria on medium, touch a loopful of
water onto the edge of a colony and let motile bacteria
swim into it.
 Then transfer loopful of water on a slide to get a faintly
turbid suspension and cover with a cover slip.
 The bacterial suspension is thus prepare with a
minimum of agitation , which would detach the
flagella.
 After 5-10 min. when many bacteria have attach to the
surface of the slide and cover-slip , apply two drops of
Ryu’s stain to the edge of cover slip and leave the stain
to diffuse in to film.
 Examine with microscope after 5-15 min.

49.  Leifson flagella stain
Solution A:
Sodium chloride 1.5 g
 Distilled water 100 ml
Solution B:
Tannic acid 3.0 g
 Distilled water 100 ml
 Solution C:
Pararosaniline acetate 0.9 g
Paraosaniline hydrochloride 0.3 g
Ethanol, 95% 100 ml
Mix equal volumes of solutions A and B; then add 2 volumes of the mixture to 1
volume of solution C. The resulting solution may be kept refrigerated for 1 to 2
months.

50.  For staining from liquid cultures, Leifson recommends
two rounds of centrifugation and final suspension in
distilled water to remove any medium components.
Place 100 ml of the liquid culture in a micro centrifuge
tube, centrifuge, and remove spent medium.
Resuspend in 100 ml of distilled water by gently
vortexing, again centrifuge, and remove supernatant.
Form a slightly cloudy emulsion by resuspending in
~200 ml of distilled water. Gently vortex. Again,
emulsions should be only slightly cloudy prior to
proceeding to staining. Optimization of the washing
procedure will most likely be necessary to maximize
quality of flagella stain.

51.  Leifson flagella stain
1. Take a prepared slide and using a wax pencil draw a
rectangle around the dried sample. Place slide on staining rack.
2. Flood Leifson dye solution on the slide within the
confines of the wax lines. Incubate at room temperature for 7 to
15 minutes. The best time for a particular preparation will
require trial and error.
3. As soon as a golden film develops on the dye surface and a
precipitate appears throughout the sample, as determined by
illumination under the slide, remove the stain by floating off the
film with gently flowing tap water. Air dry.
4. View using oil immersion, at 1,000x magnification, by
bright-field microscopy. Bacterial bodies and flagella will stain
red.

52.  When a bacterial culture is stained with Liefson stain, the
tannic acid component of the stain produce a colloidal
precipitate which can be taken up by the bacterial flagella
will become colorized which can be easily visualized using
microscopy. The concentration of the tannic acid and dye
is important in staining the bacterial flagella while the
alcohol concentration in the Liefsons stains helps in
maintaining the solubility of the components. On
microscopic observation, the bacterial cells and flagella
will stain red and the flagellar arrangement can be
visualized easily. The age of the bacterial culture,
condition of staining solutions, concentrations of the
staining solution etc can also affect the staining reaction.


53. STAINS FOR SPIROCHAETES
 Fontana’s stain for films and levaditi stain for sections.
 Large spirochaetes such as borrelia stain by ordinary
method ,including gram’s stain(giving a negative
reaction)
 But smaller ones such as treponemes and leptosira are
too thin to be seen , can be best observe by dark
ground microscope, where their bright appearance
and motility draw attention to them.

54. Fontana’s method for films:
Solutions
Fixatives:
Acetic acid 1ml
Formalin(40%) 2ml
Distilled water 100ml
Mordant:
Phenol 1gm
Tannic acid 5gm
Distilled water 100ml
Ammoniated silver nitrate
Add 10% ammonia to 0.5% solution of silver nitrate in
distilled water.

55.  Procedure:
 Treat the film three times,30 seconds each times with
fixatives.
 Wash off the fixatives with absolute alcohol and allow the
alcohol to act for 3 min.
 Pour on the mordant ,heating till steam rises, and allow it
to act for 30 seconds.
 Wash well with distilled water and dry again.
 Treat with ammoniated silver nitrate , heating till steam
rises, for 30 seconds , when the film becomes brown in
color.
 Wash well in distilled water , dry and mount in Canada
balsam as some immersion oils causes the films to fade at
once.
 Spirochetes are stained brownish black on brownish yellow
back ground.

57.  Levaditi’s method for tissue sections:
 Fix the tissue , which must be in small pieces 1 mm thick ,in
10%formaline for 24 hr.
 Wash the tissue for 1 hr in water and there after in 96-98%
alcohol for 24 hr.
 Place the tissue in 1% solution of silver nitrate for 2 hr at room
temperature, and there after at 50 degree C for 4-6 hr.
 Then rapidly wash in 10% pyridine solution.
 Transfer to the reducing fluid which contain
 4% formalin 100 part
 Acetone 10 part
 Pyridine 15 part ,which are added
immediately before use.
 Keep in this solution for 2 days at room temperature in dark.
 After washing well , dehydrate the tissue with alcohol embed in
paraffin.
 Mount it in Canada balsam.

59. Silver impregnation
 Use to see the spirochetes in tissue section where dark field microscopy not
possible , and when they are not in sufficient concentration.
 Warthin-Starry, Dieterle, Steiner silver impregnation method are used. They all
perform equivalently.
 It is also useful for flagellar demonstration.
 Reagent A
 100 ml distilled water
 5 g tannic acid,
 1.5 g ferric chloride,
 2.0 ml formalin
 1.0 ml 1 % sodium hydroxide.
 Reagent B, ammoniated silver nitrate solution:
 100 ml of 2% silver nitrate . About 10 ml of this volume is removed and saved;
to the remaining 90 ml, ammonium hydroxide is added drop wise until the
heavy precipitate that is formed is dissolved. From the 10ml previously
removed, 2% silver nitrate is added drop wise until a slight clouding appears
and persists. At this point, the pH is adjusted to 10.0 with the ammonium
hydroxide .
 Reagent B is relatively unstable, and must be used within 4 hr of preparation

60.  The smears are covered by reagent A for 2 to 4 min;
they are then rinsed in distilled water.
. After the water rinse, reagent B (pH 10.0) is added for
about 30 sec.. The smears are immediately washed
with distilled water, air-dried, and examined under oil
immersion.
 Leptospires are stained dark-brown to black .

62. CAPSULAR STAINS
 Positive Capsule Staining

 Since capsule is water soluble in nature, it is too difficult to stain the capsule with normal staining
methods. The positive capsule staining method (Anthony Method) uses two reagents to stain the
capsular material. The primary stain Crystal violet is applied over a non heat fixed bacterial
smear so that both the bacterial cells and capsular material take up the color of the primary stain.
The ionic nature of the bacterial cell binds the crystal violet stain more strongly while the non-
ionic nature of the capsule get adhere with the crystal violet stain. When the decolorizing agent
copper sulfate is added over the bacterial smear, the loosely adhered crystal violet stain is washed
off from the capsular material without removing the tightly bound crystal violet from the cell wall.
The capsular material absorbs the light blue color of the copper sulfate in contrast to the
purple bacterial cell.

 Negative Capsule Staining

 Another simple method to visualize the bacterial capsules is by using negative staining Technique.
During staining the non heat fixed bacterial smear with the acidic stains such as Nigrosin will not
penetrate the bacterial cells (since both acidic stain and bacterial surface has negative charge).
Instead the acidic stain deposits around the bacterial cells and create a dark back ground and the
bacteria appear as unstained with a clear area around them, capsule.

Note: If you heat fix the bacterial smear for capsule staining, the cells will shrink creating a hallow
zone around the bacterial cell and will be mistaken for the capsule.

64. India ink preparation:
 Mainly use for Cryptococcus neoformans, because of its
large polysaccharide capsule, can be visualized by the India
stain. Organisms that possess a polysaccharide capsule
exhibit a halo around the cell against the black background
created by the India.
 Method of Use: Mix the specimen with a small drop of
India on a clean glass slide. Place a cover slip over the
smear and press gently. The preparation should be
brownish, not black. Using reduced examine the smear
microscopically (100X) for the presence of encapsulated
cells as indicated by clear zones surrounding the cells.

65.  LIMITATIONS
 It is recommended that biochemical and/or serological
tests be performed on colonies from pure culture for
complete identification.
 The diagnosis of C. neoformans by negative staining should
be considered a presumptive result. Leukocytes, fat
droplets, and tissue cells are sometimes confused with C.
neoformans cells. Leukocytes and tissue cells may be
dissolved by adding a drop of 10% KOH.
 Some strains of C. neoformans, as well as other cryptococci
may not produce discernible capsules in vitro.

67.  The modified technique employs 2% chromium mercury. A
 small drop of CSF is placed on a clean glass slide. Then, a small
 drop of 2% chromium mercury is mixed with the CSF on the
 slide. Immediately, a small amount of India ink is added. Finally,
the cover slip is mounted and the
 preparation is observed with a bright-field microscope .
 Thus, three layers from the outer capsule that have previously
been discerned only by electron microscopy can be distinguished,
namely, the lucent stratum of the capsule, the fibrillar
material of the capsule, and the light zone .

69. Congo red stain
Materials
 Congo Red stain
 Acid fuchsin stain
 Acid alcohol
 Congo Red Capsule Stain Procedure
 1. Place a loop-full of Congo Red on a slide
 2. Mix a small amount of your organism into the drop of Congo Red.
 Spread the organism/dye suspension well on the slide
 3. Let the slide thoroughly air dry.
 4. Fix the dried slide with acid alcohol for 15 seconds.
 5. Rinse with distilled water and cover the slide with acid fuchsin for 1-5
minutes.
 6. Rinse with water and allow to air dry.
 7. Examine the slide under oil immersion.
 Cells stain red/pink, and the capsules appear as colorless halos against a
dark red background

70. Congo red

71. STAINS FOR CHAMYDIA
C. trachomatis is an obligate intracellular pathogen ,that
causes
 urethritis, proctitis,trachoma ,Infertility
 prostatitis and epididymitis in men
 cervicitis, pelvic inflammatory disease (PID), ectopic
pregnancy.
The characteristic inclusions are H P bodies(
Halberstaedter prowazek bodies) .
Staining method to these bodies are:
Giemsa
Castaneda
machhiavello

72. Giemsa stain for Chlamydia

73.  Machiavello stain:
 Reagents:
 1% Methylene Blue 1.0 gm
 Distilled water 100ML
 shelf life: 3 months storage: room temperature
 0.25% Basic Fuchsin
 0.25 gm Basic Fuchsin
 100 ml Distilled water (H2O)
 shelf life: 1 month storage: room temperature
 5% Citric Acid Solution
 0.5 gm Citric Acid (H3C6H5O7)
 95 ml Distilled water (H2O)
 shelf life: 2 months storage: room temperature

74.  Procedure:
 1. Deparaffinize and hydrate to distilled water.
 2. Stain in 1% methylene blue solution overnight.
 3. Decolorize in 95% alcohol.
 4. Wash quickly in distilled water.
 5. Counter stain in 0.25% basic fuchsin solution for 30
minutes.
 6. Decolorize rapidly in 0.5% citric acid solution for 1 or
2 seconds, never more than 3 seconds.
 7. Differentiate rapidly in absolute alcohol.
 8. Dehydrate in 95% alcohol and absolute alcohol 3 changes
each. Clear in xylene, 3 changes.
 9. Mount with Permount.
 Results: Chlamydia bright red Nuclei blue

75. Castaneda stain
 Castaneda's staining solution
Solution A
Potassium dihydrogen phosphate, 1 g
Disodium hydrogen phosphate 25 g
Distilled water 1000 ml
Formalin 1 ml
Dissolve the potassium dihydrogen phosphate in 100
ml distilled water and the disodium hydrogen phosphate in
900 ml distilled water. Mix the two solutions to give a
buffer pH 7.5, and add formaldehyde as a preservative.

76. 
Solution B
Methylene blue 1 g
Methanol 100 ml
Staining solution
Solution A 20 ml
Solution B 0.15 ml
Formalin 1 ml
Safranine-acetic acid
Safranine (0.2% aqueous solution) 1 part
Acetic acid (0.1% aqueous solution) 3 parts

77.  Procedure.
* Prepare films from infected tissue and dry in air
* Apply the stain for 3 min.
* Drain, do not wash
* Counter stain for a 1-2 seconds in safranin-
acetic acid
* Wash in running water, blotdry.
 Elementary bodies : blue. Cell nuclei and cytoplasm:
red.

79.  SUMMARY:
 Special stain are used to see the structures of cell which
are not able to stain by routine methods.
 Also helpful for provisional diagnosis of disease.
 Fluoresce method will increase the sensitivity and time
saving method.
 Also useful to proceed further before any expensive
diagnosis method is used .

80. REFERENCES
 Monica Cheesbrough
 Koneman
 Ananthanarayan
 Mackie & McCartney

81. THANK YOU