Vol. 53,No. 6 Rinlcd in 11.S. A.
NOTES ON TECHNIC
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
ELIMINATION OF STATIC ELECTRICITY DURING PARAFFIN SECTIONING
STEVEN R. SCADDING, Department of Zoology, University of Guelph, Guelph, Ontario, Canada, NIG 2 WI The problem of static electricity interfering with paraffin sectioning is widespread and particularly acute for those of us doing serial sectioning in older buildings where the humidity is very low in winter. The main source of the problem is the friction of the knife as it cuts the parafin block and the accumulation of a static charge on the ribbon. The main static charge creating a problem is in most cases that charge which accumulates on the ribbon itselfcausing it to adhere to anything nearby, including the microtome or the technician’s hands. The traditional remedies such as breathing on the block and knife or boiling water next to the microtome (Humason 1972) are often ineffective. In addition, the latter suggestion poses a significant burn hazard to the technician. Other suggestions for elimination of static electricity, appear to have limited usefulness, e.g. that of Dichiara and Koopmans (1977) requires a constant source of dry ice and a room of moderate humidity. The suggestion of Bryan and Hughes (1976) that the technician be electrically grounded exposes the technician to the hazard of potentially fatal electrical shock if contact with a live electrical source is made. Mattheij and Dignum (1975) compared the merits of radioactive polonium-210 devices, which discharge static electricity on the ribbon by emission of ionizing alpha particles, versus electrostatic ionizing devices and concluded that the poloniuni-2 10 device posed a considerably greater hazard to personnel in the laboratory. However. the electrostatic ionizing apparatus they used must be constructed specifically for this application. We have found that a standard Simco Static Eliminator, although not intended for use in microtomy, is a safe and simple means of applying this principal to the discharge of the static electricity on the parafin ribbon. The device consists of a Simco Midget Power Unit (Model D167RY-$82.20) and a Shockless One-Point Static Bar ($48.25) manufactured by The Simco Co., 920 Walnut Street, Lansdale, Pa., 19446. The ionizing device waq mounted on a gooseneck arni (from an old magnifier-Edmund Scientific. No. 70,241) so that it could be moved into position about 2 cm from the forming paraffin ribbon when needed (Fig. 1) and removed when not in use. This proved more convenient in our laboratory than permanent mounting on the microtome. However, any convenient method of mounting the static bar would work since the only constraint is that the discharge point must not be more than about 2 to 3 crn from the paraffin ribbon for the ribbon 10 be discharged. The microtome was grounded to the power unit. Electrostatic deionizers of this type are quite safe (Mattheij and Dignum 1975) and even touching the discharge point produces almost no detectable sensation of 35 1
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
352
STAIN TECHNOLOGY
FIG. 1. The static bar ahown in p i t i o n during acctioning. It is not necessary to have the power unit near the microtome. In use it mats fcyeral feet away but is placed beside the microtome hem for pucp0”c” of illustration.
shock. Since this device will operate for many years, the higher initial cost compares favorably with the cost of polonium strips ($30.00-$40.00) which must be replaced every 12 to 18 months. Since four Static Bars can be operated from one power unit, static elimination by this method will be much cheaper than polonium strips where more than one microtome is in use. In our laboratory this device has effectively eliminated the static electricity problem and allows safe and convenient serial sectioning even on the driest days. REFERENCES
Bryan,J. H. D. and Hughes, R. L. 1976. A simple and inexpensive static eliminator for paraffin sectioning. Stain Technol. 50: 397-398. Dichiara, J. and Koopmans, H. 1977. A new method for mducing static electricity during paraffin sectioning. Stain Technol. 52: 353-354. Humason, G. L. 1972. Animal Ticnu Techniques. W.H. F m m a n and Co., San Francisco. Mattheij, J. A. M.and Dignum, P.H. 1975. A device for the complete elimination of static electricity in parafin sectioning. Stain Technol. 50: 157-159.
NOTES O N TECHNIC
353
THEUSE OF GLYCERIN TO IMPROVE MORPHOLOGY OF TISSUES FROZENSECTIONED FOR ULTRASTRUCTURAL CYTOCHEMISTRY M. PINO,'LOUISTERRACIO' AND PATRICK W. BANKSTON, Departmi .f RICHARD
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
Anatomy, Hahncmann Medical College and Hospital, Phila&&hia, Pemuylvania 19102 To facilitate reagent penetration tissues prepared for ultrastructural enzyme cytochemistry are usually reacted as 20-200 pm sections. For sectioning, the freezing microtome has been replaced in many laboratories by the TC-2 Tissue Sectioner (Du Pont Instruments, Sorvall Operations, Newton, Connecticut) or the Vibratome (Ted Pella Co., Tustin, California) both of which are capable of slicing unfrozen tissue into sections of suitable thickness without undue tissue damage. However, frozen sections are still widely used. Frozen sections of fixed material prepared for enzyme cytochemistry often exhibit suboptimal submicroscopic morphology because it is often necessary to decrease fixation times or reduce fixative concentrations to limit enzyme inhibition. We find that this is especially evident in frozen sections of delicate fetal tissues. We now present a simple method which maintains excellent morphology during frozen sectioning. Tissues are fixed, diced, and rinsed in buffer according to the established protocol for the enzyme to be localized. After the buffer wash, tissue pieces are immersed for 10-15 minutes in 20% glycerin in 0.1 M sodium cacodylate buffer, pH 7.4, at 4 C and then frozen on the stage of a freezing microtome in the same solution. Due to the cryoprotective properties of glycerin, upon cooling the tissues become firm enough to be cut, yet not hard enough to fragment. Structural damage due to ice crystal formation is minimized. Sectioning can be rapid with additional chilling of the mount as necessary. Many tissue blocks can be cut simultaneously; often, complete cross sections of tissue blocks will slide onto the knife's surface intact. After collection in an appropriate buffer the sections are reacted in cytochemical media and processed further by routine electron microscopic methods. This technique has been used successfully to localize endogenous peroxidase (Fig. I), acid phosphatase (Fig. 2), catalase, and thiamine pyrophosphatase activities in fetal and adult tissues. REFERENCES Graham, R. C. and Karnovsky, M. J. 1966. The early stages of absorption of injected horseradish peroxidase in the pmximal tubules of mouse kidney; ultrastructural cytochemistry by a new technique. J. Histochem. Cytochem. 14: 291 -302. Novikoff, A. B. 1963. Lysmomes in the physiology and pathology of cells: contributions of staining methods. Ciba Foundation Symposium on Lysosornes. A. V. S. de Reuck and M. P. Cameron, eds. J. A. Churchill Ltd., London, pp. 3 6 7 3 .
' Present address: Kresge Eye Institute, Wayne State University, Detroit, Michigan 48201.
* Present address: W. Alton Jones Cell Science Center, Lake Placid, New York 12946.
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
!jTAIN TECHNOLOGY
FIG. I. Endogenous peroxidase in Kupffer cell from a 15-day fetal rat liver. Fixation: 30 minutes in 1.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, p l l 7.2; reacted 2 hours according to the hydrogen peroxide. ER, endoplasmic reticulum; method of Graham and Karnovsky (1966) using 0.01'%1
m. mitochondria; N, nucleus of Kupffer cell; NE, nuclear envelope. Lead citrate stain. X 28,500. FIG. 2. Liwr of 19-day fetal rat reacted for acid phosphatase. Fixation: I hour in 2'7opformaldehyde2% glutaraldehydc in 0.1 M sodium cacodylatc buffer, pH 7.2; reacted 20 minutes according 10 the method of Novikoff (1963). H, nucleus of hepatocyte; KC, Kupffer cell; L, sinusoidal lumen; gly, glycogen lakes; m, mitochondria; PE, phagocytized erythrocyte nucleus; arrows, acid phosphatase positivc phagosoma. Lead citratc stain. X 4,460.
NOTES ON TECtiNIC
355
DOUBLE FIXATION A N D ACETOCARMINE STAINING FOR PERMANENT CHROMOSOMAL PREPARATIONS OF ALGAL FLAGELLATES
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
R. SHYAM, Algae Laboratory, National Botanical Research Institute, Lucknow 226001, India The chromosomes of unicellular or colonial algal flagellates are very difficult to stain and to make permanent preparations of. The present technique is based on double fixation followed by staining with acetocarmine. Proccdure. Two fixatives are used. Fixative I is prepared as outlined by Nissenbaum (1953) and consists of 10 ml saturated mercuric chloride, 2 ml glacial acetic acid, 2 ml formalin and 5 ml tertiary butyl alcohol. Fixative I1 is Carnoy’s 1:3 acetic-alcohol as applied by Cave and Pocock (1951); however, in the present case the glacial acetic acid was supersaturated with ferric acetate by boiling over 10 g of this salt for 45 to 60 min in a round-bottom flask using a reflux condenser. After standing for a fortnight, 1 part of the supernatant is combined with 3 parts of absolute alcohol to make the fixative. Killing and fixation are begun immediately on removal of the organisms from culture. A drop of actively dividing organisms is placed on a slide, a drop of freshly prepared Fixative I is added and the two are thoroughly mixed with a stainless steel needle. As soon as the fluid evaporates the slide is transferred to a Coplin jar containing freshly prepared fixative I1 and left for 4-24 hr. Next the slide is rinsed with acetic acid-alcohol (1:3) and placed in a Coplin jar containing saturated acetocarmine stain and left for 15 min. A second transfer to fresh acetocarmine gives better staining. The slide is then removed from the Coplin jar and wiped free of stain except around the specimens. A drop of fresh acetocarmine is put on the slide and a coverslip put in place. T h e slide is then warmed gently over a low spirit lamp flame and the specimens flattened with thumb pressure. Excess acetocarmine is removed with a filter paper after which the slide is ready for microscopic examination. T h e pressure required for proper chromosome separation may vary. Slides are made permanent, first, by removing the coverslip in 45% acetic acid and completing dehydration in two changes of absolute alcohol, 5 min each. Alcohol is removed from the preparations gradually using a series of three intermediate mixtures (3:1, 1: 1, 1:3) between absolute alcohol and xylene. T h e coverslip is replaced using Canada balsam. Results and Discussion. Nucleoli and flagella stain lightly but distinctly. Chromosomes stain a deep crimson, sometimes blackish red (Fig. 1). T h e cytoplasm stains faintly without interfering with the staining of nucleus or chromosomes. The procedure reported here is highly advantageous in securing (a) rapid and firm adhesion and fixation of the flagellates on the slide itself thereby avoiding the difficulties usually experienced in repeated centrifugation to bring down the cells for fixation, mordanting, washing and staining (cf: Godward 1966), (b) better contrast in chromosome staining and (c) effective spreading of chromosomes. Preparations mounted permanently by the above method have remained unchanged for 10 years. Originally applied to unicellular and colonial Volvocales,
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
356
STAIN TECHNOLOGY
Ro. 1. A well-spread metaphase plate of Gaium $ectorah.
X
2OOO.
the method has been found, with slight modifications to suit individual groups, to be equally good for algal flagellates belonging to the Euglenophyceae and Dinphyceae. REFERENCES Cave, M. S. and PomcL, M. A. 1951. The acetocarmine technique applied to the colonial Volvocals. Stain. Technol. 26: 173-174. Codward, M. B. E. 1966. TXc Chromosomes of A l p . Arnold, London. Nisstnbaum,C. 1953. A combined method for the rapid fixation and adhesion ofciliates and flagellates. Science 118: 3 12-3 13.
A COLORFAST STAIN FOR POLYESTEREMBEDDED LIGNEOUS TISSUES D. T. BUCKBURN, Department of Botany, UniversiQ of Adelaide, South Australia, 5001 A clearing and mounting technique described by Christophel and Blackburn (1975) has proven useful in the preparation of a permanent cleared leaf collection. A problem associated with the long term storage of the leaves has become evident however. It has been found that the safranin 0 used for staining the preparations fades with time. Fading is accelerated by exposure to light and appears to be related to the presence of traces of catalyst in the mountant. The mounting medium is a low wax, high clarity polyester resin. Any clear potting resin such as Biopot may be used. Its polymerization is catalyzed by methyl ethyl ketone peroxide.
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
NOTES O N TECHNIC
357
Catalysis is effected by the generation of free radicals from the methyl ethyl ketone peroxide. Presence of these groups even in hardened resin accelerates light induced decomposition of the organic stain molecules. To overcome this problem a n inorganic stain has been tested and found to be superior in all respects to safranin. Johansen (1940) described a test for lignin which used potassium permanganate as a n intermediate reagent (Maule test). Cleared leaves stained only in 0.1 M neutral aqueous potassium permanganate gave excellent results for selective staining of venation only, resulting in a high degree of contrast between veins and niesophyll (Fig. 1). T h e pigment itself is manganese dioxide, deposited from the reduction of the permanganate in the presence of lignin. This compound is chemically stable and there is no reason to believe that it will fade with time. T h e stain is fine grained and not discernible by optical microscopy at loo0 times magnification. T h e new modified version of the procedure described by Christophel and Blackburn (1975) is as follows. 1. Soak either fresh or dried leaves in 15% w/w potassium hydroxide until they are decolorized, changing the solution every two days as necessary. T h e most satisfactory containers for this have been found to be glass Petri dishes, with the leaves placed in them under discs of plastic mesh screening to prevent floating. The time for decolorization varies considerably with the specimens and may be as long as six weeks. In the case of stubborn leaves, gentle warming may facilitate clearing. 2. Wash the leaves under a gentle stream of water for two minutes and transfer to saturated aqueous chloral hydrate until totally clear. This should take from two to seven days with slight warming being useful on slow material.
PI
NOTES ON TECHNIC
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
ELIMINATION OF STATIC ELECTRICITY DURING PARAFFIN SECTIONING
STEVEN R. SCADDING, Department of Zoology, University of Guelph, Guelph, Ontario, Canada, NIG 2 WI The problem of static electricity interfering with paraffin sectioning is widespread and particularly acute for those of us doing serial sectioning in older buildings where the humidity is very low in winter. The main source of the problem is the friction of the knife as it cuts the parafin block and the accumulation of a static charge on the ribbon. The main static charge creating a problem is in most cases that charge which accumulates on the ribbon itselfcausing it to adhere to anything nearby, including the microtome or the technician’s hands. The traditional remedies such as breathing on the block and knife or boiling water next to the microtome (Humason 1972) are often ineffective. In addition, the latter suggestion poses a significant burn hazard to the technician. Other suggestions for elimination of static electricity, appear to have limited usefulness, e.g. that of Dichiara and Koopmans (1977) requires a constant source of dry ice and a room of moderate humidity. The suggestion of Bryan and Hughes (1976) that the technician be electrically grounded exposes the technician to the hazard of potentially fatal electrical shock if contact with a live electrical source is made. Mattheij and Dignum (1975) compared the merits of radioactive polonium-210 devices, which discharge static electricity on the ribbon by emission of ionizing alpha particles, versus electrostatic ionizing devices and concluded that the poloniuni-2 10 device posed a considerably greater hazard to personnel in the laboratory. However. the electrostatic ionizing apparatus they used must be constructed specifically for this application. We have found that a standard Simco Static Eliminator, although not intended for use in microtomy, is a safe and simple means of applying this principal to the discharge of the static electricity on the parafin ribbon. The device consists of a Simco Midget Power Unit (Model D167RY-$82.20) and a Shockless One-Point Static Bar ($48.25) manufactured by The Simco Co., 920 Walnut Street, Lansdale, Pa., 19446. The ionizing device waq mounted on a gooseneck arni (from an old magnifier-Edmund Scientific. No. 70,241) so that it could be moved into position about 2 cm from the forming paraffin ribbon when needed (Fig. 1) and removed when not in use. This proved more convenient in our laboratory than permanent mounting on the microtome. However, any convenient method of mounting the static bar would work since the only constraint is that the discharge point must not be more than about 2 to 3 crn from the paraffin ribbon for the ribbon 10 be discharged. The microtome was grounded to the power unit. Electrostatic deionizers of this type are quite safe (Mattheij and Dignum 1975) and even touching the discharge point produces almost no detectable sensation of 35 1
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
352
STAIN TECHNOLOGY
FIG. 1. The static bar ahown in p i t i o n during acctioning. It is not necessary to have the power unit near the microtome. In use it mats fcyeral feet away but is placed beside the microtome hem for pucp0”c” of illustration.
shock. Since this device will operate for many years, the higher initial cost compares favorably with the cost of polonium strips ($30.00-$40.00) which must be replaced every 12 to 18 months. Since four Static Bars can be operated from one power unit, static elimination by this method will be much cheaper than polonium strips where more than one microtome is in use. In our laboratory this device has effectively eliminated the static electricity problem and allows safe and convenient serial sectioning even on the driest days. REFERENCES
Bryan,J. H. D. and Hughes, R. L. 1976. A simple and inexpensive static eliminator for paraffin sectioning. Stain Technol. 50: 397-398. Dichiara, J. and Koopmans, H. 1977. A new method for mducing static electricity during paraffin sectioning. Stain Technol. 52: 353-354. Humason, G. L. 1972. Animal Ticnu Techniques. W.H. F m m a n and Co., San Francisco. Mattheij, J. A. M.and Dignum, P.H. 1975. A device for the complete elimination of static electricity in parafin sectioning. Stain Technol. 50: 157-159.
NOTES O N TECHNIC
353
THEUSE OF GLYCERIN TO IMPROVE MORPHOLOGY OF TISSUES FROZENSECTIONED FOR ULTRASTRUCTURAL CYTOCHEMISTRY M. PINO,'LOUISTERRACIO' AND PATRICK W. BANKSTON, Departmi .f RICHARD
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
Anatomy, Hahncmann Medical College and Hospital, Phila&&hia, Pemuylvania 19102 To facilitate reagent penetration tissues prepared for ultrastructural enzyme cytochemistry are usually reacted as 20-200 pm sections. For sectioning, the freezing microtome has been replaced in many laboratories by the TC-2 Tissue Sectioner (Du Pont Instruments, Sorvall Operations, Newton, Connecticut) or the Vibratome (Ted Pella Co., Tustin, California) both of which are capable of slicing unfrozen tissue into sections of suitable thickness without undue tissue damage. However, frozen sections are still widely used. Frozen sections of fixed material prepared for enzyme cytochemistry often exhibit suboptimal submicroscopic morphology because it is often necessary to decrease fixation times or reduce fixative concentrations to limit enzyme inhibition. We find that this is especially evident in frozen sections of delicate fetal tissues. We now present a simple method which maintains excellent morphology during frozen sectioning. Tissues are fixed, diced, and rinsed in buffer according to the established protocol for the enzyme to be localized. After the buffer wash, tissue pieces are immersed for 10-15 minutes in 20% glycerin in 0.1 M sodium cacodylate buffer, pH 7.4, at 4 C and then frozen on the stage of a freezing microtome in the same solution. Due to the cryoprotective properties of glycerin, upon cooling the tissues become firm enough to be cut, yet not hard enough to fragment. Structural damage due to ice crystal formation is minimized. Sectioning can be rapid with additional chilling of the mount as necessary. Many tissue blocks can be cut simultaneously; often, complete cross sections of tissue blocks will slide onto the knife's surface intact. After collection in an appropriate buffer the sections are reacted in cytochemical media and processed further by routine electron microscopic methods. This technique has been used successfully to localize endogenous peroxidase (Fig. I), acid phosphatase (Fig. 2), catalase, and thiamine pyrophosphatase activities in fetal and adult tissues. REFERENCES Graham, R. C. and Karnovsky, M. J. 1966. The early stages of absorption of injected horseradish peroxidase in the pmximal tubules of mouse kidney; ultrastructural cytochemistry by a new technique. J. Histochem. Cytochem. 14: 291 -302. Novikoff, A. B. 1963. Lysmomes in the physiology and pathology of cells: contributions of staining methods. Ciba Foundation Symposium on Lysosornes. A. V. S. de Reuck and M. P. Cameron, eds. J. A. Churchill Ltd., London, pp. 3 6 7 3 .
' Present address: Kresge Eye Institute, Wayne State University, Detroit, Michigan 48201.
* Present address: W. Alton Jones Cell Science Center, Lake Placid, New York 12946.
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
!jTAIN TECHNOLOGY
FIG. I. Endogenous peroxidase in Kupffer cell from a 15-day fetal rat liver. Fixation: 30 minutes in 1.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, p l l 7.2; reacted 2 hours according to the hydrogen peroxide. ER, endoplasmic reticulum; method of Graham and Karnovsky (1966) using 0.01'%1
m. mitochondria; N, nucleus of Kupffer cell; NE, nuclear envelope. Lead citrate stain. X 28,500. FIG. 2. Liwr of 19-day fetal rat reacted for acid phosphatase. Fixation: I hour in 2'7opformaldehyde2% glutaraldehydc in 0.1 M sodium cacodylatc buffer, pH 7.2; reacted 20 minutes according 10 the method of Novikoff (1963). H, nucleus of hepatocyte; KC, Kupffer cell; L, sinusoidal lumen; gly, glycogen lakes; m, mitochondria; PE, phagocytized erythrocyte nucleus; arrows, acid phosphatase positivc phagosoma. Lead citratc stain. X 4,460.
NOTES ON TECtiNIC
355
DOUBLE FIXATION A N D ACETOCARMINE STAINING FOR PERMANENT CHROMOSOMAL PREPARATIONS OF ALGAL FLAGELLATES
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
R. SHYAM, Algae Laboratory, National Botanical Research Institute, Lucknow 226001, India The chromosomes of unicellular or colonial algal flagellates are very difficult to stain and to make permanent preparations of. The present technique is based on double fixation followed by staining with acetocarmine. Proccdure. Two fixatives are used. Fixative I is prepared as outlined by Nissenbaum (1953) and consists of 10 ml saturated mercuric chloride, 2 ml glacial acetic acid, 2 ml formalin and 5 ml tertiary butyl alcohol. Fixative I1 is Carnoy’s 1:3 acetic-alcohol as applied by Cave and Pocock (1951); however, in the present case the glacial acetic acid was supersaturated with ferric acetate by boiling over 10 g of this salt for 45 to 60 min in a round-bottom flask using a reflux condenser. After standing for a fortnight, 1 part of the supernatant is combined with 3 parts of absolute alcohol to make the fixative. Killing and fixation are begun immediately on removal of the organisms from culture. A drop of actively dividing organisms is placed on a slide, a drop of freshly prepared Fixative I is added and the two are thoroughly mixed with a stainless steel needle. As soon as the fluid evaporates the slide is transferred to a Coplin jar containing freshly prepared fixative I1 and left for 4-24 hr. Next the slide is rinsed with acetic acid-alcohol (1:3) and placed in a Coplin jar containing saturated acetocarmine stain and left for 15 min. A second transfer to fresh acetocarmine gives better staining. The slide is then removed from the Coplin jar and wiped free of stain except around the specimens. A drop of fresh acetocarmine is put on the slide and a coverslip put in place. T h e slide is then warmed gently over a low spirit lamp flame and the specimens flattened with thumb pressure. Excess acetocarmine is removed with a filter paper after which the slide is ready for microscopic examination. T h e pressure required for proper chromosome separation may vary. Slides are made permanent, first, by removing the coverslip in 45% acetic acid and completing dehydration in two changes of absolute alcohol, 5 min each. Alcohol is removed from the preparations gradually using a series of three intermediate mixtures (3:1, 1: 1, 1:3) between absolute alcohol and xylene. T h e coverslip is replaced using Canada balsam. Results and Discussion. Nucleoli and flagella stain lightly but distinctly. Chromosomes stain a deep crimson, sometimes blackish red (Fig. 1). T h e cytoplasm stains faintly without interfering with the staining of nucleus or chromosomes. The procedure reported here is highly advantageous in securing (a) rapid and firm adhesion and fixation of the flagellates on the slide itself thereby avoiding the difficulties usually experienced in repeated centrifugation to bring down the cells for fixation, mordanting, washing and staining (cf: Godward 1966), (b) better contrast in chromosome staining and (c) effective spreading of chromosomes. Preparations mounted permanently by the above method have remained unchanged for 10 years. Originally applied to unicellular and colonial Volvocales,
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
356
STAIN TECHNOLOGY
Ro. 1. A well-spread metaphase plate of Gaium $ectorah.
X
2OOO.
the method has been found, with slight modifications to suit individual groups, to be equally good for algal flagellates belonging to the Euglenophyceae and Dinphyceae. REFERENCES Cave, M. S. and PomcL, M. A. 1951. The acetocarmine technique applied to the colonial Volvocals. Stain. Technol. 26: 173-174. Codward, M. B. E. 1966. TXc Chromosomes of A l p . Arnold, London. Nisstnbaum,C. 1953. A combined method for the rapid fixation and adhesion ofciliates and flagellates. Science 118: 3 12-3 13.
A COLORFAST STAIN FOR POLYESTEREMBEDDED LIGNEOUS TISSUES D. T. BUCKBURN, Department of Botany, UniversiQ of Adelaide, South Australia, 5001 A clearing and mounting technique described by Christophel and Blackburn (1975) has proven useful in the preparation of a permanent cleared leaf collection. A problem associated with the long term storage of the leaves has become evident however. It has been found that the safranin 0 used for staining the preparations fades with time. Fading is accelerated by exposure to light and appears to be related to the presence of traces of catalyst in the mountant. The mounting medium is a low wax, high clarity polyester resin. Any clear potting resin such as Biopot may be used. Its polymerization is catalyzed by methyl ethyl ketone peroxide.
Biotech Histochem Downloaded from informahealthcare.com by Chinese University of Hong Kong on 12/27/14 For personal use only.
NOTES O N TECHNIC
357
Catalysis is effected by the generation of free radicals from the methyl ethyl ketone peroxide. Presence of these groups even in hardened resin accelerates light induced decomposition of the organic stain molecules. To overcome this problem a n inorganic stain has been tested and found to be superior in all respects to safranin. Johansen (1940) described a test for lignin which used potassium permanganate as a n intermediate reagent (Maule test). Cleared leaves stained only in 0.1 M neutral aqueous potassium permanganate gave excellent results for selective staining of venation only, resulting in a high degree of contrast between veins and niesophyll (Fig. 1). T h e pigment itself is manganese dioxide, deposited from the reduction of the permanganate in the presence of lignin. This compound is chemically stable and there is no reason to believe that it will fade with time. T h e stain is fine grained and not discernible by optical microscopy at loo0 times magnification. T h e new modified version of the procedure described by Christophel and Blackburn (1975) is as follows. 1. Soak either fresh or dried leaves in 15% w/w potassium hydroxide until they are decolorized, changing the solution every two days as necessary. T h e most satisfactory containers for this have been found to be glass Petri dishes, with the leaves placed in them under discs of plastic mesh screening to prevent floating. The time for decolorization varies considerably with the specimens and may be as long as six weeks. In the case of stubborn leaves, gentle warming may facilitate clearing. 2. Wash the leaves under a gentle stream of water for two minutes and transfer to saturated aqueous chloral hydrate until totally clear. This should take from two to seven days with slight warming being useful on slow material.
PI
