The Peracetic-Orcein-Halmi Stain: A Stain for Connective Tissues

A selective stain useful for the study of connective tissues is described. The stain demonstrates elastic and oxytalan fibers as well as fibrils in mucous connective tissues previously undescribed. Reticular fibers are not stained. The stain may be used on sections that have been fresh frozen or fixed in formalin or ethanol. Sections are deparaffinized, washed in absolute ethanol, oxidized in peracetic acid 30 min, washed in running water, stained in Taenzer-Unna orcein 15 min, 37°C, differentiated in 70% ethanol, washed in running water, stained in Lillie-Mayer alum hematoxylin 4 min, blued in running water, and counterstained 20 sec in a modified Halmi mixture of 100 ml distilled water, 0.2 gm light green SF, 1.0 gm orange G, 0.5 gm phosphotungstic acid and 1.0 ml glacial acetic acid. Sections are rinsed briefly in 0.2% acetic acid in 95% ethanol, dehydrated and mounted.     

A Tetrachrome Stain for Fresh, Mineralized Bone Sections, Useful in the Diagnosis of Bone Diseases

Fresh, unprocessed bone is ground to sections 75-100 μ thick, stained in an aqueous solution composed of fast green FCF, 0.1 gm; orange G, 2.0 gm; distilled water, 100.0 ml; and adjusted to pH 6.65, then in a mixture of 1 part alcoholic solution of 0.25% celestine blue B and 9 parts of alcoholic solution of 0.1% basic fuchsin. Surface stain is removed by grinding sections to 50 μ and washing them in 1% invert soap (Zephiran) to remove adherent debris. (Commercial detergents and alkaline soaps may interfere with chromophore groups of the dyes.) Wash in tap water; rinse in distilled water and differentiate in 1% acetic alcohol. Dehydrate in ascending alcohols, clear in xylene and mount permanently in a neutral, synthetic resin. Active osteoid seams stain dark to light green; resting osteoid seams, red to bright orange red; transitional osteoid seams, geenish-yellow, orange red to red; older, partly mineralized matrix, orange; new, partly mineralized matrix, red; osteocyte nuclei, red; osteoblasts and osteoclasts, greenish-blue to dark purple nuclei and green or light green cytoplasm. Hyper-trophic and differentiating cartilage cells are stained light pink and dark red respectively. The staining reactions are consistent; the solutions are stable.     

Oil Blue Na as A Stain for Rubber in Sectioned or Ground Plant Tissues

Oil blue NA (Calco), a stain which colors rubber bright blue, has been used effectively in studying the distribution of rubber in several plant species. Fresh or fixed sections are cut, bleached with Javelle water or NaOCl solution, treated with 9% KOH in 95% ethanol, washed with several changes of water and finally with 95% ethanol, and stained with 0.05% oil blue NA in 70% ethanol. Sections are rinsed in 50%' ethanol, placed in 40% glycerin, and mounted in glycerin jelly.

For the detection of changes in the distribution and character of rubber in milled or ground tissues, much the same staining procedure is followed. The stained tissues usually are examined and dissected under a stereoscopic microscope, a procedure which permits rubber to be recognized by both its staining reaction and by a more specific property, elastic elongation.

A microscopic technic is presented whereby it is possible to determine approximately the relative proportion of dispersed and coagulated rubber latex in unstained tissues.     

The Use of Bismarck Brown Y in some new Stain Schedules

The staining quality of Bismarck brown Y may be improved and sterility maintained by adding 5% phenol to a 1% aqueous solution. Use the phenolic Bismarck brown in combination with iron alum hematoxylin except for stripped epidermis in the following procedures:

Stem and Root Schedule: Mordant sections from water in 4% iron alum for 10 minutes. Rinse in distilled water and stain in 0.5% aqueous hematoxylin for 1 minute or until darkly stained. Rinse in distilled water and destain in 2% iron alum until a gray color appears. Rinse thoroly in distilled water and intensify hematoxylin by transferring sections to 0.5% aqueous lithium carbonate until the desired black color appears. Rinse thoroly in distilled water and stain for 1-5 minutes in phenolic Bismarck brown. Rinse in distilled water, dehydrate successively in 30, 50, 70, 95 and 100% alcohol. Clear in methyl salicylate for 5 minutes, then to xylene for 3-5 minutes, and mount in balsam.

Middle Lamellae in Wood: Destain more thoroly in 2% iron alum than for the general stem and root schedule, and intensify in lithium carbonate for a longer period (about 1 hour).

White Potato Tuber Sections: Modify above schedule by reducing time of destaining in 2% iron alum to about 30-60 seconds and intensify hematoxylin until starch grains appear bluish in color. Stain in phenolic Bismarck brown for 1-2 minutes.

Wheat Grain Sections: Fix grain for sectioning when in “dough” stage. Use schedule the same as for potato tuber except for reducing time of staining in phenolic Bismarck brown to about 45 seconds.

Tradescantia zebrina Epidermis: Strip epidermis from leaf while submerged in water. Fix in 100% alcohol 10 minutes, pass thru 95, 70, 50, 30, and 10% alcohol to water. Stain in phenolic Bismarck brown for 10-20 minutes. Dehydrate, clear in methyl salicylate and mount in balsam.     

Modified Whipf's Polychrome: A Connective Tissue Stain with Special Application for Demonstrating Leishmania

A polychrome stain procedure was developed to demonstrate amastigotes of the protozoan parasite Leishmania braziliensis as well as cytoplasmic and other tissue components in cutaneous lesions of infected animals. The procedure is as follows: stain nuclei for 10 minutes with an iron hematoxylin containing 0.5% hematoxylin and 0.75% ferric ammonium sulfate dissolved in 1:1 0.6 N H₂SO₄:95% ethanol; rinse 4 minutes in distilled water. Cytoplasmic staining is achieved by exposing tissues for 10 minutes to a solution containing 0.25% Biebrich scarlet, 0.45% orange G, 0.5% phosphomolybdic acid and 0.5% phosphotungstic acid in 1% aqueous acetic acid. These first two solutions are modified from Whipf's polychrome stain. Sections are differentiated for 10 seconds in 50% ethanol, rinsed in water, stained 3 minutes in 0.1% aniline blue WS in saturated aqueous picric acid, rinsed in water and differentiated for 1 minute in absolute ethanol containing 0.05% acetic acid. Mordanting overnight in 6% picric acid in 95% ethanol produced optimal results.

This procedure was applied to sectioned material from experimental animals with various protozoa. Trypanosoma cruzi, Besnoitia Jellisoni, Toxoplasma gondii and especially Leishmania braziliensis were well demonstrated. Combining cytoplasmic dyes and phosphomolybdic-phosphotungstic acids into one solution afforded differential staining of tissues by Biebrich scarlet and orange G; connective tissues were stained by this solution. Substantially improved definition of connective tissues resulted after counterstaining. This procedure differs from the Massou sequence in which connective tissues are first stained by cytoplasmic dyes along with other tissues and then destained prior to specific counter-staining. in comparing dyes structurally related to Biebrich scarlet, it was found that Crocein scarlet MOO, but not Poncenu S, was an acceptable substitute. Sirius supra blue GL and Sirius red FSBA were not useful as replacements for aniline blue WS in this procedure.     

Differential in Toto Staining of Bone, Cartilage and Soft Tissues

The following method for staining bone and cartilage allows study of the gross cleared specimen and does not injure the tissues for subsequent microscopic study: Fix in 10% neutral formalin; bleach thoroughly in 3% H₂O₂ in sunlight. Wash in distilled water. Stain bone 24 hours in 0.01 g. of Biebrich scarlet in 100 ml. of distilled water. Destain in 95% alcohol until soft tissues and cartilage are colorless. Stain cartilage 24 hours in a pH2 buffer solution of 2.1g. of citric acid per 100 ml. of water with 0.001 g. of methylene blue. Destain in pH2 buffer solution until soft tissues are pale. Dehydrate in two changes of 95% alcohol in preparation for clearing. (This completes the destaining and may remove too much stain from the cartilage if destaining in the pH2 solution has been carried too far.) Place in Groat's clearing fluid and cover loosely so that the alcohol may evaporate, or remove the alcohol in vacuo. Groat's Mixture No. 19 is usually satisfactory.

For a combined stain, first stain bone, as above, and then apply the cartilage stain.

Seal jars with an ordinary liquid wood glue such as LePage's.     

Mallory's Connective Tissue Stain Following Hematoxylin

The following combination of hematoxylin with Mallory's connective tissue stain is useful in bringing out nuclei as well as in differentiating tissue:

Slightly overstain in Mayer's hematoxylin (50 g. potassium alum and 0.2 g. sodium iodate added to 1 liter 0.1% aqueous hematoxylin). Wash; and stain 30 seconds to 1 minute in 0.04% aqueous acid fuchsin-Stain 4 minutes in: 0.5 g. anilin blue and 2 g. orange G dissolved hi 100 cc. of 1% aqueous phosphomolybdic acid. Pass thru 95% alcohol to absolute; clear in xylol and mount in balsam.     

Staining Aspirated Human Bone Marrow with Domestic Wright Stain

The method employs the domestic Wright stain for the staining of aspirated human bone marrow. Freshly distilled water, pH 6.0 to 6.4, is used. Wright stain, 0.5 cc, is placed upon the air-dried preparation and permitted to act for two minutes. The stain is then diluted with 2 cc. distilled water and permitted to act for from 5 to 10 minutes. After washing off the stain with distilled water, the preparation is placed into a decolorizer (acetone 0.5 cc, pure methyl alcohol 5.0 cc, and 100 cc. distilled water, pH 6.0 to 6.4) for differentiation from 1 to 5 seconds, rinsed, washed under running water and permitted to air-dry. A well stained and differentiated preparation shows the “Romanovsky effect”, and the sharpness of minute structures obtained compares favorably with control preparations stained with German dyes.

The bone marrow should be prepared as described. The Wright stain marketed by the National Aniline and Chemical Co., N. Y. was found to be reliable as regards staining quality of registered batches. One photomicrograph, showing bone marrow cells from pernicious anemia, is included.     

Staining Raw and Cooked Apple Tissues for Study of Cell Wall Structure

Permanent preparations were made of paraffin sections from raw and cooked apple tissues stained with microchemical color reagents for pectins and pentosans. Sections stained with ruthenium red to show pectins were dehydrated and covered in balsam, and sections stained with diphenylene diamine acetate (DDA) to show pentosans were washed with water and covered in Clearcol.

Cooking was accomplished by steaming cubed histological samples. Both raw and steamed specimens were fixed in FAA in a vacuum chamber, dehydrated and cleared in tertiary butyl alcohol, and embedded in paraffin. Paraffin sections first fixed to slides with Haupt's adhesive were further stabilized by immersing in a 1% celloidin solution after dissolving the paraffin.

Ruthenium oxychloride flakes were dissolved in a Coplin jar of water containing 2 drops of ammonium hydroxide. Rehydrated sections were stained in ruthenium red 30 minutes and rinsed in water. Three methods of further preparation follow: (1) Flood sections with 10% gum arabic; drain and air-dry thoroughly; immerse in xylene 5 minutes; cover in balsam. (2) Drain and air-dry sections; if desired, counterstain dry sections with Johansen's fast green solution; immerse in xylene; cover in balsam. (3) Dehydrate by dipping in 70%, 95%, and absolute ethyl alcohol; immerse in xylene; cover in balsam.

DDA was made by heating 15 g. of benzidine in 150 ml. of glacial acetic acid and 450 ml. of water until dissolved, then adding water to make 750 ml. of solution. Rehydrated sections were stained 4 hours in DDA, washed, stained 5 minutes in Congo red (Congo red, 5 g.; NaOH, 5 g.; water, 100 ml.), washed, and covered in Clearcol.

An Autotechnicon was used for dehydration, clearing, infiltration, deparaffinizing sections, and staining. Procedures that necessarily remained manual were fixation in a vacuum chamber, and all operations that followed staining.

Ruthenium red, though the best available indicator for pectins, may not be specific for these substances. DDA and ruthenium red stained identical structures in hypodermis and cortex. DDA also stained cuticle, hence was more useful than ruthenium red for delineating that portion. DDA sections were better for photomicrography, and for measuring thickness of cell walls. Neither stain prevented the study of cell walls in polarized light.     

Further Experiments with the Masson Trichrome Modification of Mallory's Connective Tissue Stain

Several dyes, notably ponceau 2R, azofuchsin 3B, nitrazine yellow, and Biebrich scarlet may replace imported “ponceau de xylidin” in the Masson ponceau acid fuchsin mixture. Of these Biebrich scarlet appears to be the best and may be used without acid fuchsin.

A mixture of equal parts of 5% solutions of phosphomolybdic and phosphotungstic acids is much superior to either acid alone and gives adequate mordanting in 1 minute at 22°C.

With the fast green modification, times in plasma and fiber stains can be reduced to 2 minutes each. With anilin blue a 4-minute plasma stain is required. One-minute final differentiation in 1% acetic acid is adequate.

Primary mordanting of formalin material may be accomplished by 5 minutes in saturated aqueous mercuric chloride or 2 minutes in saturated alcoholic picric acid. Three minutes washing in running water is required after these mordants.     

A Durable Nissl Stain for Frozen and Paraffin Sections

Frozen sections, 25-50 /j. thick, of formalin-fixed nervous tissues are mounted following the Albrecht gelatin technic. Paraffin sections, 15 p., are deparaffinized and transferred to absolute ethanol. The slides are then coated with celloidin. Both frozen and paraffin sections subsequently follow the same steps: absolute ethanol-chloroform (equal parts) for at least 20 min, 95% ethanol, 70% ethanol (1-3 min), then rinsed in distilled water. Sections are stained in Cresylechtviolett (Chroma) 0.5% aqueous solution containing 4 drops of glacial acetic acid per 100 ml, rinsed in distilled water, agitated in 70% ethanol until excess stain leaves the slide, and rinsed in 95% ethanol. Sections are then dehydrated in absolute ethanol, followed by butanol, cleared in xylene, and enclosed in permount.     

Brilliant Cresyl Blue as a Stain for Plant Chromosomes

Methods are proposed for staining plant chromosomes with the dye brilliant cresyl blue, and for making these stained preparations permanent by using polyvinyl alcohol mounting medium.

The stain, which is composed of 2% brilliant cresyl blue in 45% aqueous acetic or propionic acid, is used with fixed material in making smear preparations. The technics for staining are similar to those employed in the aceto-carmine method.

The mounting medium is made by mixing 56% polyvinyl alcohol, which is diluted in water to the consistency of thick molasses, with 22% lactic acid and 22% phenol by volume. The permanent slides are made by floating off the cover slip of the temporary slide in 70% alcohol, then applying the mounting medium and replacing the cover slip.

The chief advantages of the methods described are:

1)The preparation of the stain is rapid and simple. The batch of stain will be good with the first try.

2)The staining procedure in some instances is shorter than when using aceto-carmine.

3)The stain shows a high degree of specificity for nuclear structures and gives better results than aceto-carmine when used on certain plant tissues.

4)A minimum number of cells is lost in making the slides permanent when using polyvinyl alcohol mounting medium as the slide and cover slip are run through only one solution prior to mounting.

5)The mounting medium dries rapidly and this shortens the time required before critical examination of the permanent mounts can be made.     

Stain Technic for Nissl's Granules Following Alcohol-Formol Fixation

1. Nerve tissue is fixed 2-4 hrs in a 5% solution of strong formalin in commerical 95% alcohol.

2. If dehydration is perfect, either chloroform or xylol may be used as a clearing agent.

3. A slow method of paraffin infiltration is advisable.

4. Sections should be cut 10-12 microns in thickness.

5. Coplin staining jars should be annealed by placing them on a rack in a pan of cold water, bringing the water to the boiling point, and allowing the jars to stand in boiling water for twenty minutes.

6. One per cent aqueous solutions of either methylene blue or Grübler's Neutral Roth are used as specific stains for Nissl's granules.

7. These stains are heated to boiling in a beaker, the slides are placed in the Coplin jars which are partially submerged in boiling water, and the hot stain poured into the jars. The flame beneath the water bath is turned down and the slides left for 20 minutes.

8. The excess of primary stain is washed off in 25% and 50% alcohol and the slides passed rapidly thru the alcohol series to absolute alcohol, and finally to xylol.

9. When counterstaining is desired, nigrosin in 1% aqueous solution, methyl orange, saturated solution in 50% alcohol, or, a 0.5% solution of eosin in 50% alcohol are recommended. These stains are used cold, and the slides are merely dipped in them after the excess of primary stain has been washed out in 25% and 50% alcohol.

10. If a cold primary stain is desired, a saturated solution of thionin in distilled water, acidified with 1% carbolic acid, will prove specific for the Nissl substance. Sections should be stained 5–10 minutes in thionin, then passed rapidly thru to absolute alcohol, and xylol. The same counterstains may be used as in the hot method.

11. Sections prepared by the hot method show little tendency to fade after ten years use.

12. Excepting neutral red, all the stains used in this technic are carried by the National Aniline and Chemical Company and are satisfactory. Coleman and Bell neutral red may be substituted for Grübler's Neutral Roth with good results.     

A Versatile New Mineralized Bone Stain for Simultaneous Assessment of Tetracycline and Osteoid Seams

A versatile mineralized bone stain (MIBS) for demonstrating osteoid seams and tetracycline fluorescence simultaneously in thin or thick undecalcified sections has been developed. Bone specimens are fixed in 70% ethanol, but 10% buffered formalin is permissible. Depending upon one's preference, these specimens can be left unstained or be prestained before plastic embedding. Osteoid seams are stained green to jade green, or light to dark purple. Mineralized bone matrix is unstained or green. Osteoblast and osteoclast nuclei are light to dark purple, cytoplasm varies from slightly gray to pink. The identification of osteoid seams by this method agrees closely with identification by in vivo tetracycline uptake using the same section from the same biopsy. The method demonstrates halo volumes, an abnormal, lacunar, low density bone around viable osteocytes in purple. This phenomenon is commonly seen in vitamin D-resistant rickets, fluorosis, renal osteodystrophy, hyperparathyroidism, and is sometimes seen in fluoride treated osteoporotic patients. In osteomalacic bone, most osteoid seams are irregularly stained as indicated by the presence of unmineralized osteoid between mineralized lamellae. The method has been used effectively in staining new bone formation in hydroxyapatite implants and bone grafts. Old, unstained, plastic embedded undecalcified sections are stained as well as fresh sections after removal of the coverslip. This stain also promises to be valuable in the study of different metabolic bone diseases from the point of view of remodeling, histomorphometry, and pathology.     

A Controllable Differential Stain for the Hypophysis by Adaptation of the Mallory Triple Stain

A selective and controllable staining method for the hypophysis has been developed with rat material, using Mallory's triple stain as a basis.

Fix in Zenker neutral formol for 6 hours. Longer fixation is undesirable. Transfer to 30% alcohol plus a few drops of a saturated solution of I₂ in aqueous KI over night. Gradually complete dehydration and clear in cedar oil. Infiltrate with a paraffin mixture (paraffin, rubber-paraffin, bayberry wax and beeswax). Section 3-Sμ. Hydrate to distilled water, placing a few drops of a KI-I₂ solution in the 50% alcohol. Stain in 1% acid fuchsia for 30 minutes. Rinse, and differentiate in a weak NH₄OH solution (one drop 28% NH₄OH to 200 cc. HOH). When differentiation is complete, transfer to a 0.5% phosphomolybdic acid solution for 3 minutes, after first stopping the differentiation with a 0.1% HC1 solution and then rinsing with distilled water. Stain for one hour in a solution of: 1% anilin blue, water soluble, 2% orange 6, and 1% phosphomolybdic acid. Rinse in distilled water plus a few cubic centimeters of the stain. Differentiate in 95% alcohol, transfer to absolute alcohol and clear in a mixture of 30% oil of cedar, 40% oil of thyme, 15% absolute alcohol and 15% xylene. Finally, transfer to xylene and mount.     

A Versatile Stain for Pollen Fungi, Yeast and Bacteria

A versatile stain has been developed for demonstrating pollen, fungal hyphae and spores, bacteria and yeasts. The mixture is made by compounding in the following order: ethanol, 20 ml; 1% malachite green in 95% ethanol, 2 ml; distilled water, 50 ml; glycerol, 40 ml; acid fuchsin 1% in distilled water, 10 ml; phenol, 5 g and lactic acid, 1-6 ml. A solution has also been formulated to destain overstained pollen mounts. Ideally, aborted pollen grains are stained green and nonaborted ones crimson red. Fungal hyphae and spores take a bluish purple color and host tissues green. Fungi, bacteria and yeasts are stained purple to red. The concentration of lactic acid in the stain mixture plays an important role in the differential staining of pollen. For staining fungi, bacteria and yeasts, the stain has to be acidic, but its concentration is not critical except for bacteria. In the case of pollen, staining can be done in a drop of stain on a slide or in a few drops of stain in a vial. Pollen stained in the vial can be used immediately or stored for later use. Staining is hastened by lightly flaming the slides or by storing at 55±2 C for 24 hr. Bacteria and yeasts are fixed on the slide in the usual manner and then stained. The stock solution is durable, the staining mixture is very stable and the color of the mounted specimens does not fade on prolonged storage. Slides are semipermanent and it is not necessary to ring the coverslip provided 1-2 drops of stain are added if air bubbles appear below the coverslip. The use of differentially stained pollen mounts in image analyzers for automatic counting and recording of aborted and nonaborted pollen is also discussed.     

A Tribasic Stain for Thin Sections of Plastic-Embedded, OsO4-Fixed Tissues

Thin (0.5-1 μ) sections of plastic-embedded, OsO₄-fixed tissues were attached to glass slides by heating to 70 C for 1 min. A saturated solution combining toluidine blue and malachite green was prepared in ethanol (8% of each dye) or water (4% of each dye). Methacrylate or epoxy sections were stained in the ethanol solution for 2-5 min. The water solution was more effective for some epoxy sections (10-80 min). Epoxy sections could be mordanted by 2% KMnO₄, in acetone (1 min) before use of the aqueous dye, reducing staining time to 5-10 min and improving contrast. Aqueous basic fuchsin (4%) was used as the counter-stain in all cases; staining time varied from 1-30 min depending upon the embedding medium and desired effects, methacrylate sections requiring the least time. In the completed stain, nuclei were blue to violet; erythrocytes and mitochondria, green; collagen and elastic tissue, magenta; and much and cartilage, bright cherry red. Sections were coated with an acrylic resin spray and examined or photographed with an oil-immersion lens.     

Metallic Lakes of the oxazines (Gallamin Blue, Gallocyanin and Coelestin Blue) as Nuclear Stain Substitutes for Hematoxylin

Becher's investigations upon the soluble metallic lakes of the oxazines have been re-investigated, extended and results described. Gallamin blue, gallocyanin and coelestin blue in combination with ferric ammonium sulfate gave the best results. The dyes are dissolved in a five per cent aqueous solution of ferric ammonium sulfate. The solution is boiled for 2-3 minutes, cooled, filtered and ready for immediate use. The iron lakes of these dyes stain nuclei excellently giving a deep blue or blue black in 3-5 minutes. No differentiation with acid is required. Coelestin blue gives the most stable solution and is recommended as a routine nuclear stain. The protoplasm remains practically colorless and counter-staining with acid dyes such as ethyl-eosin, orange G, or fuchsin gives pictures which cannot be distinguished from a good hematoxylin stain.

Counter-staining with van Gieson solution is also possible. Benda's modification of the van Gieson solution is recommended. Staining of fat with Sudan, scarlet red, etc., does not interfere with nuclear staining by these dyes.

As applied to the central nervous system these dyes are far superior to hematoxylin. Ganglion and glia cells are as excellently stained as with thionin.

The most widely used fixatives, namely formaldehyde, Mueller-formaldehyde, Zenker's and alcohol, give equally as good results. The nature of the staining process is briefly discussed and a prospectus offered.     

A Differential Stain for Rubber in Guayule

The following schedule, which combines an intense blue stain for rubber with sharply contrasting red counterstains, has been found satisfactory for use in an anatomical study of rubber deposition in guayule: Cut fresh or fixed sections about 50 to 100 % thick, transfer to 50% ethanol. Extract with acetone 5 minutes, treat with 1% NaOCl 5 minutes, saponify with 10% KOH in 95% ethanol 15 minutes, rinse 3 times with 50% ethanol, stain in oil blue NA (Calco) with safranin and Congo red 30 minutes at 55° C. Rinse in 50% ethanol 2 (or more) times to remove excess stain and mount in Karo syrup.     

Fluorochrome Staining of Multilamellar Liposomes

Multilamellar liposomes can be stained with such fluorochromes as acridine orange, eosin Y, neutral red, and thiazine red. The liposomes are brought into a 1% solution of the fluorochrome; 5-10 minutes later they are centrifuged and washed by resuspending in water or phosphate buffered saline three times. The last pellet is resuspended and a drop studied with the fluorescence microscope (1000 × magnification). The fluorochrome is seen to be accumulated in the liposomal membranes.

Acridine orange could also be trapped in the aqueous compartments of the liposomes but the trapped fluorochrome was gradually lost from the liposomes. Part of the fluorochrome, however, remained associated with the liposomal membranes for a long time.

Additional experiments justify the conclusion that an equilibrium is maintained between fluorochromes in the aqueous and lipid phases.