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Techniques in Anatomy, Cytology and Histochemistry of Plants
Book · November 2006
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Sugarcane Breeding Institute
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Techniques in Physiology, anatomy, Cytology &
Histo-Chemistry of Plants
CONTRIBUTORS
R. KARUPPAIYAN
Dr. K. NANDINI
Scientist (Plant Breeding)
ICAR Research Complex for NEHR
Gangtok, Sikkim-737 102
Associate Professor (Physiology)
College of Horticulture
Kerala Agricultural University
Vellanikkara, Thrissur-680 656
Dr. E.V. ANOOP
Assistant Professor (Wood science)
College of Forestry
Kerala Agricultural University
Vellanikkara, Thrissur-680 656
Dr. T. GIRIJA
Assistant Professor (Physiology)
College of Horticulture
Kerala Agricultural University
Vellanikkara, Thrissur-680 656
Sh. M.ABDUL NIZAR, M.Sc. (Botany)
Sr. Scientist
NBPGR Regional Station
Akola, Maharashtra
Department of Plant Physiology
College of Horticulture
Kerala Agricultural University
Vellanikkara, Thrissur-680 656
PREFACE
This practical manual is intended for under graduate, post graduate students as well as
research scholars and teachers from Agricultural, Horticultural and Forestry Sciences.
Accordingly the exercises were fitted. This manual is divided into four parts. In the first part
general guidelines to the students and teachers to comply or equip to laboratory culture, some
basic concepts and terminologies in plant anatomy, cytology and cytochemistry, guidelines to
the use microscope, microtome, procedures for preparation of glassware cleaning solutions,
stains, fixatives, etc were dealt with.The second part of the manual was devoted to impart skill
oriented anatomical techniques like sectioning, smearing, peeling, staining, mounting, etc. In
addition we tried to impart practical knowledge on anatomy of monocot, dicot plant organs
with typical examples.Cytological techniques like understanding chromosomal behaviour during
mitosis, meiosis, ascertaining pollen fertility, etc were given in third parts; And the last part
was allotted to practical histochemistry (cytochemistry). There were 12 practical neatly
described to understand the localization of phytochemicals like carbohydrate, protein, lipids,
nucleic acids, phenol, lignin, mucilage, cutin, suberin, etc.
This book is a self-explanatory manual with neat drawings, photographs. Suitable
illustrations and examples were given at eligible place. Keeping in mind the non- availability
and prohibitory cost of some chemicals or instruments / apparatus and also to satisfy the thrust
of many researcher alternative procedures wherever available were given. The specialty in the
manual is that it contains both old and new methods and practices. It is versatile and suitable
for from beginner to researcher. Each exercise has tips on dos and don’ts. In any one interested
to learn more details about any of the exercises we suggest the following information sources.
Some information was drawn from these sources.
Vellanikkara
25-11-2006
K. Nandini
R. Karuppaiyan
CONTENTS
Practical
No
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
Title
Laboratory etiquette
Basic concepts and terminology
Proper use of a microscope
Measurement of microscopical objects and drawings
Understanding photo-micrographic camera system
Preparation of glassware cleaning solutions
Preparation of materials for anatomical studies
Understanding some anatomical techniques
(Maceration, Peeling and Smearing)
How to make a hand section
Mounting and ringing
Study on microtome
Pre-treatment for microtome sectioning
Taking section in microtome
Staining techniques
Study the structure of a typical plant cell
Anatomy of monocot and dicot plant organs
Examining anatomy of monocot root-Maize
Examining anatomy of dicot root-Bengal gram
Examining anatomy of monocot stem-Maize
Examining anatomy of dicot stem-Cucurbita
Examining anatomy of monocot leaf-Maize
Examining anatomy of dicot leaf-Mango
Ecological anatomy and special anatomical features
Study on stomata in hydrophyte and xerophyte
Study on mitosis in root tips
Study on meiosis in floral buds
Study on pollen fertility
Cytochemistry & stain specificity
Localisation of cellulose /mucilage/cutin/suberin
Localisation of total insoluble polysaccharides
Localisation of cellulose
Localisation of reducing substances
Localisation of pectic acid/lignin/tannin
Localisation of lignin
Localisation of phenols
Localisation of cutin and suberin
Localisation of protein
Localisation of Lipids
Localisation of nucleic acids (DAN and RNA)
Suggested Readings
Page No
1-5
6-11
12-18
19-20
21-23
24
25-28
29-31
32-34
35-36
37-39
40-42
43-49
50-56
57
58-61
62
63
64
65
66
67
68
69
70-72
73-75
76
77-78
79-80
81
82-83
84
85-86
87
88
89
90-91
92-93
94-95
96
Practical 1
LABORATORY ETIQUETTE
Inside the laboratory one is expected to maintain the dignity and decorum, for it
allows the work to be done properly. Some of the instruction to and expectation from the
students / researchers are given below for perusal.
General instructions
1.
We want you to be a trust-worthy researcher
2.
You are requested to strictly follow the instruction of your teacher
3.
Whenever you feel any difficulty never hesitate to consult your teacher
4.
Work with sincerity, confidence, faith, courage and dedications.
5.
Confine to your own work as far as possible
6.
You must be well conversant with theoretical parts of items of which you
have to perform the practical
7.
You must record in your lab note book the sequence of treatments,
observation, protocols, results, or any events. Nothing should be left to
memory.
8.
Laboratory notes should have your own identity or originality.
9.
Any observation or drawing whether morphology or anatomy must be faithful
and you must record or draw what you see. Do not imagine anything and do
not let others to say your results are cooked or fag.
10.
Those who are routinely working in the laboratory are requested to write in the
indent register for chemicals before it is exhausted. Never resort to post
mortem purchase exercise. This will unnecessarily delay your work
11.
Use hand gloves, lab coat and shoes while working in the lab
12.
Every students in the laboratory is an investigator and he or she must perfect
(complete) his own exercise or technique thoroughly and also budget his
own timings.
13.
Close the doors while you leave the lab and hand over the keys to security /
concerned person. Do not carry keys with you.
14.
Since laboratory shall be open during night (but access restricted), we
encourage you working during night as well. However, don’t be lonely in
the lab. Do your work along with your class mates. Do not forget to bring
drinking water and torch light / candle with match box during night.
15.
Never borrow or lend any instrument without informing to the teachers.
16.
Do not play rough with your pointed instruments otherwise their points will be
spoiled; your body will also be damaged.
1
17.
Concentrate only on your work; do not make noise; better if you observe silent
while working in the lab and do not disturb others.
This is more important
1. While leaving the laboratory kindly see or ensure that burners, distillation units,
laminar air flow, autoclave, water taps, AC units and all electric switches
except refrigerators are switched off. Computers are properly shut downed and
plugs were disconnected to prevent from possible damage due to lightening.
2. Laboratory is your home. So keep it neat and clean. Understand ‘Cleanliness is
Godliness’ and have the habit of spending at least a hour in the lab. Keep on
practicing hard and troublesome exercises. ‘Practice makes a Man Perfect’.
3. Do not fill your mind of complaint about teachers, departments and lack of
facilities in the institution. Blame your own-self for any errors and kindly adjust
yourself to work efficiently with existing facilities.
Precaution with acids
1.
Never add water into acids-As a thumb rule acid must be poured into water
2.
Avoid close eye contact with acid fumes as it can cause irritation
Glassware and instruments
1. Keep your things, instruments, apparatuses very neat and clean
2. Arrange glassware and instruments in their original place. Arrange chemicals
as per alphabetical order
3. All glassware must be washed with detergent / cleaning solution before and
after use
4. Handle the glassware, instruments and apparatus with utmost care. Do not
break it. If broken, kindly inform to your teacher and carefully dispose off the
broken pieces.
5. Pipettes should be separate for each chemicals, so also the measuring cylinders
6. For pipeting acids and hazardous chemicals you should use pipettes with safety
bulb
2
Chemicals and First Aid
1. Chemicals and Reagents must be labeled properly before you use.
2. Do not use very old or oxidized or reduced chemicals
3. Work very carefully with poisonous and hazardous chemicals (e.g. chemicals
like HgCl, Ammonia, acids like H2SO4, mutagens like EMS)
4. Anhydrous solution and dehydrating fluids must be kept in well stopper bottle
5. KI and I2 containing bottles must never be kept in an almirah containing other
chemicals as it sublimes and all labels of bottles turn dirty black
6. Solids should not be thrown into sink. Wash the sink after chemicals are
thrown in the sink
7. Osmic acid fumes should never be inhaled or brought close to eyes
Using an electronic balance
1. Keep the balance away from fan, in plain surface, carefully adjust the platform to
rectify any tilting / vibration with the aid of mercury leveler attached in some of
the balance
2. Connect the balance preferably with UPS at the time of weighing
3. Switch on balance 2 to 5 min before weighing to make it to attain stability
4. Weighing must be done by keeping non-absorbent papers on pans (use papers
from old chemical catalogue like Merck) and then keeping the materials to be
weighed
Stains and Fixatives
1.
Balsam, Iodine Potassium iodide stains on exposure to light becomes acidic
and is then harmful for stains. Therefore the containers should be protected
from light by keeping in amber bottles or store in dark room
2.
By the method of trial and error select the best possible stains and fixatives.
3.
Never gets discouraged / frustrated if get poor results. Analyze the cause of
your failure yourself. Judgment must be based on killing, fixing, infiltration,
microtome, staining and mounting techniques. Every step in a process must
be carefully examined. Try and try again. You will attain perfection
4.
Understand the staining procedure thoroughly before you proceeds
5.
As a rule never leave tissues / specimen in for very long in killing solutions,
dehydrating fluids or paraffin bath
3
WORKS TO THE TEACHERS
Teachers should ensure accessibility and availability of the following items to the
students
1. Microscope-simple and compound-Each student should get one working
microscope
2. Microtome-Rotary, sledge and hand microtome
3. Microscopic slides: Size of a microscopic slide is 25 mm x 75 mm (3” x 1”). The
standard thickness is 1 mm; however, slightly thicker slide are also used. Cover
slips are available with different size and thickness. Squares and Circles shape
cover slips come in 15, 18, 22 and 22 mm standard sizes. Rectangles are either 22
or 24 mm wide and 30,40,50,60 and 70 mm long. The thickness is designated by
number. No. 0 to No. 3. Zero number cover slips are used for oil immersion. The
standard one is No 1 (0.17 mm) or No 2 (0.18 mm?) thickness and 18 or 22 mm
circles / 22 mm squares / 22 x 40 mm rectangles. Plastic cover slips are costly but
durable.
4. All necessary chemicals and stains
5. A razor strope and a hone / razor / budding knife-one to each student
6. Petri dishes-Several numbers are required
7. Watch glass-Three or more
8. Depression slides or culture slides or cavity slides-One box
9. Small dropping bottles for reagents-Half dozen number or more
10. Soft cloth for polishing slides and cover slips-Three Number (15 x 15 cm)
11. A tile-white glazed, 6” square, half of it must be enameled black
12. Slide box-Made of bakelite or wood-Three Numbers
13. Hand lens /pocket lens-Planetic type-10 to 20 numbers
14. Enamel tray-3 to 5
15. Wash bottle or squeeze bottle
16. Ordinary pipette with safety bulb
17. Staining trays (with cavity)- Stender dishes (Flat type) 3 or 4
18. Staining trays: Coplin jars. Vertical and flat -Ten to twenty pieces
19. Spirit lamp
20. Balsam bottle with glass rods-One number
21. Glass marking pencil
22. Camel hair brushes (No 1 & 2): -10 numbers
23. Sticker paper for labeling
4
24. Paraffin embedding oven (for microtome)
25. Scissors-Small and medium size-one each
26. Scalpel-one
27. Penknife-one
28. Needles and forceps
29. Dropper bottles-permits the stain to come out in drops-3 Nos
30. Droppers / ink pillar (long handles and short)-Two each
31. Blotting paper
32. Tissue paper
33. Filter paper
34. Pencil sharpener and Eraser (Rubber)
35. Band aid or adhesive wrap to tie around finger while taking section-1 roll
36. Surgical gloves-one per student
Note to the teachers: The students may be instructed to bring some of the above items like
needle, forceps, brush, razor, pencil, band aid wrap, gloves, etc. to economies office
spending
5
Practical 2
BASIC CONCEPTS AND TERMINOLOGY
The students / research scholars are advised to through with meaning and
terminologies of various anatomical structures. You will be in darkness if you proceed to
work without understanding the structural differences as well terminologies used to
describe them. Given below are commonly used terminologies in plant anatomical
research.
Definition
1. Anatomy: deals with gross as well as minute structure-both external and internal of
the organism
2. Cytology: Branch of biology that deals with the study of structure, function,
development, reproduction, and life history of cells
3. Cytogenetics. Branch of biology that deals with the correlated study of genetics
and cytology.
4. Histology: it is apart of anatomy and it deals with the structure of tissue
5. Histochemistry: deals with in situ localization of biochemical constituent at
cellular levels.
HOW TO DESCRIBE PLANES OF AN ANATOMICAL SECTION
Sometimes it is desired to study the materials from various regions. As such sections
are cut in different planes. The following are commonly used terminology
i). With reference to a cylindrical structure like stem,
1. Transverse section (TS): The section is cut by passing the razor’s edge right angle
to the longitudinal axis. It is also called cross section (c.s) since it is at cross (right
angle 0 tot eh length of the specimen.
2. Longitudinal section (LS): The section is cut by passing the razor’s edge at right
angle to the transverse axis. In this plane it may be cut along any of the radii, the
plane being known as RLS i.e. Radial Longitudinal Section. If it is cut along the
tangent it is termed as TLS i.e. Tangential Longitudinal Section
ii). In case of dorsi-ventral organs like leaf
Vertical Transverse section: In case of dorsi-ventral organs like leaf generally
section is cut in transverse plane and is known as Vertical Transverse section (VTS),
being cut in vertical plane.
Transverse Sections yield the greatest amount of anatomical data. However, the
other two longitudinal sections are needed for the best documentation of wood anatomy.
6
TISSUES AND TISSUE SYSTEMS
Tissue is a group of cells that are similar in origin, structure and function. Groups of
tissue constitute tissue system; groups of tissue systems constitute an organ; and a group of
organs into an organism. The general classification of tissues is – Meristematic tissues
and Permanent tissues. Meristematic tissues composed of cells that have the capacity to
divide, and give rise to permanent tissues. Apical meristem, intercalary meristem and
lateral meristem are the three types of primary meristem; lateral meristem of roots (e.g.
cambium), secondary cambium of stem and cork cambium are the example for secondary
meristem. Permanent tissues may be of primary or secondary according to their origin from
primary or secondary meristem. Examples are Parenchyma, Collenchyma, Sclerenchyma,
Xylem, Phloem, Fibres, lactiferous tissues, glandular hairs, etc
Tissue system
1. Dermal tissue system (tegumentary): Outer tissues that covers the plant e.g.
epidermis or periderm
•
Epidermis: Outer layer of cells, primary in origin, composed of different kinds
of cells. It may be provided with cuticle or different kinds of hairs. Its function
is to protect the internal tissues
•
Cuticle: A layer of waxy materials; more or less impervious to water; present
on outer wall of the epidermis
•
Stomata: Composed of guard cells and stoma (subsidiary cells). Its function is
gaseous exchange
•
Hairs /trichome: It is an epidermal appendage. Variable in size and shape;
glandular or non-glandular. The terminology used are,
1. Puberulous: Minutely present
2. Tomentose: Densely covered with short, soft, wool like hairs
3. Villouse: Thickly covered with long and soft hairs
4. Velutinous: Clothed with a velvety covering composed of erect,
straight, moderately firm hairs
5. Wooly: Densely covered with soft, long, curled hairs looking like a
wool
6. Pilose: Thinly covered with long, soft hairs
7. Scabrous: Feeling roughish or gritty to touch
8. Hispid: Beset with rigid or bristly hairs
9. Stellate: Star like hairs having radiating branches; hairs once or twice
forked are often treated as stellate
10. Hirsute: provided with rather rough and course hairs
7
11. Strigose: With sharp, apprised straight hairs, stiff and often basally
swollen
12. Sericeous: Provided with silky hairs
2.
Ground tissue system / Periblem: The entire complex of ground tissues i.e. tissues
other than epidermis or periderm and vascular tissues. Hypodermis, cortex,
endodermis, Pericycle and pith or pith rays constitute the periblem
i.
Cortex: The ground tissue region between the vascular system and the
epidermis; a primary tissue region
ii.
Hypodermis: Layer or layer s of cells just beneath the epidermis; referred in
case it is morphologically distinct from other cortical layers.
iii.
Aerenchyma: A parenchyma tissue; characterized by large intercellular
spaces
iv.
Bulliform cell: An enlarged epidermal cell in the leaves of grasses. Also
known as motor cell
v.
Chlorenchyma: A parenchyma tissue containing numerous chloroplasts. e.g.
leaf mesophyll
vi.
Collenchyma: Supporting tissue, composed of more or less living cells; walls
unevenly thickened
vii.
Idioblast: A special cell in the tissue which differs in form, size or contents
from other cells in the same tissue
viii.
Parenchyma: A tissue composed of parenchymatous cell. Parenchyma cell
contains nucleate protoplast; thin walled; varied in size and shape. Palisade
parenchyma-characterized by elongated cells and their arrangement
perpendicular to the surface of the leaf. e.g. leaf mesophyll parenchyma.
Spongy parenchyma is also a leaf mesophyll parenchyma characterized by
large intercellular spaces; varying cell shapes
ix.
Sclerenchyma: A tissue composed of sclerenchyma cells. e.g. fibre, sclereids,
stone cells, etc
x.
Endodermis: A layer of ground tissue surrounding the vascular region.
Specific wall characters-casparian strips or secondary thickening; generally
innermost layer of the cortex in stems and roots of seed plants
xi.
Pericycle: A part of the ground tissue of the stele located between the phloem
and endodermis; present in root and absent in most of the stems.
xii.
Rays: Medullary rays and pith rays-they are strips of parenchyma between the
vascular bundles
3. Vascular tissue/ Plerome: They were differentiated from procambium during
primary growth.
i. Cambium: A lateral meristem which differentiates from its parallelly arranged
derivatives into vascular tissue or tissues of the cork
8
ii. Phloem: The major food conducting tissue of the vascular plants; composed of
sieve elements (sieve tube, sieve plate and companion cells), fibres and
sclereids. Based on position of phloem three types can be distinguishedExternal phloem if primary phloem situated outside the primary xylem;
included phloem or interxylary phloem-if secondary phloem embedded in
the secondary xylem of certain dicotyledons and included phloem or
intraxylary phloem if primary phloem located internally to the primary xylem.
Based on size and differentiation the classification of phloem is Primary
phloem (differentiated from procambium during primary growth; first
protophloem is formed followed by metaphloem) and Secondary phloem-The
phloem tissue formed by the vascular cambium during secondary growth of the
plant
iii. Xylem: The tissue mainly responsible for conduction of water; characterized
by the presence of trachiary elements; also serves as supporting tissue. Based
on size and differentiation the classification of xylem are-Primary XylemXylem tissue differentiated from procambium during primary growth; grouped
into protoxylem and metaxylem. Protoxylem-The first formed elements of the
xylem in the plant; generally smaller in size as compared to metaxylem;
Metaxylem-differentiated after the protoxylem; Secondary xylem-The xylem
tissue formed by vascular cambium during secondary growth of vascular plant.
On the basis of its position the classification is 1)Endarch-Protoxylem (oldest
element) is closest to the centre of the axis. e.g. stem and leaves; 2) ExarchXylem strand where protoxylem are farthest from the centre of the axis e.g.
root; 3) Mesarch-Xylem strand where protoxylem are in the centre. In roots
the primary xylem may present with two protoxylem poles (diarch), three
(triarch), four (tetrarch), more than four (polyarch).
VASCULAR BUNDLE: Xylem and phloem constitute the plerome or vascular bundle.
Sometimes sclerenchyma or thick walled parenchyma associated with VB and appear
like a cap on the phloem or xylem side in cross section. this is called Bundle cap,
while Bundle sheath refers to layer or layers of cells enclosing a VB; composed of
either parenchyma or sclerenchyma. Three types of vascular bundle-1)Radial: when
xylem and phloem form separate groups and lie on different radii alternating with
each other; typical of root; 2)Concentric-One of the vascular bundle (xylem or
phloem) occupies the centre and is surrounded by another tissue; 3) Conjoint-A
vascular bundle in which both xylem and phloem lie one above the other. there are
two sub types viz., collateral VB: Phloem is present only on one side of the xylem
and bicollateral VB where phloem is present on both internal and external faces of
the xylem.
VB in relation to cambium: 1) Open VB: When cambium is present in between xylem
and phloem tissues of conjoint, collateral or bicollateral VB; 2) Closed VB:-When
cambium is absent from VB as in monocotyledons
STELE: The central cylinder of the axis; portion of the plant axis consisting of the
vascular system and its associated ground tissues (i.e. Pericycle, pith, interfascicular
region). The classification is-1) Protostele: Simple type of stele; composed of central
9
solid core of xylem surrounded by phloem; 2) Siphonostele-A type of stele where
central part is occupied by pith; a hollow cylindrical vascular tissue
GROWTH
Primary growth: The growth of the successively formed roots, vegetative and
reproductive shoots, from the time of their origin from apical meristem until the
complete formation of organs; begins from apical meristem; results in derivatives
meristem, protoderms, ground tissues and procambium and subsequently into older
tissues
Secondary growth: Conspicuous due to increase in the thickness of root and stem; results
due to the formation of secondary vascular cambium; supplemented by the activity of
the phellogen (cork cambium) producing periderm in the cortical region.
SECONDARY GROWTH IN THE CORTICAL REGIONS
Bark: A non-technical term; all the tissues outside the vascular cambium or the xylem;
divided into outer dead bark and inner living bark
Periderm: Secondary protective tissues derived from the phellogen (cork cambium);
composed of cork (phellum), cork cambium (phellogen) and secondary cortex
(phelloderm)
Phellum (cork): Protective tissue; consist of non-living dead cells with suberised walls;
replaces epidermis.
Phelloderm(secondary cortex): A tissue formed by phellogen on the opposite side of the
cork i.e. towards the inner side; similar in appearance to cortrical parenchyma
Lenticel: Special formation in the periderm; isolated area with suberised or non suberised
cells; showing numerous inter cellular spaces between them.
SECONDARY GROWTH IN THE CORTICAL REGIONS
Fascicular cambium: Vascular cambium originating from procambium; the first formed
cambium in the vascular bundles
Interfascicular cambium: Vascular cambium originating between vascular bundles or
fascicles; in the interfascicular parenchyma
Interfascicular region: Tissues region located between vascular bundles in the stem; also
known as rays or medullary.
WOOD-SECONDARY XYLEM
Annual ring: A growth layer of xylem; successive concentric rings formed year after year
10
Spring wood (Early wood): Secondary xylem formed during spring; large and thin
elements forming a distinct concentric ring
Autumn wood (Late wood): Secondary xylem formed during autumn; small and thick
elements forming a distinct concentric ring
Heart wood: The inner layer of wood which in a growth has stopped to function; without
living cells and reserve materials; dark in colour
Sap wood: The outer layers of wood which are functioning which living cells, which
contain reserve food materials; lighter in colour.
Soft wood: Wood produced by coniferous plants
Hard wood: Wood produced by dicotyledons
Porous wood: Secondary xylem (wood) with vessels
SPECIAL CELLS / ANOMALOUS STRUCTURES
Tylosis: In wood; Outgrowth from a ray or axial parenchyma cells through a pit cavity in
vessel wall; blocks of lumen of the vessel partially or completely.
Interxylary phloem (Internal Phloem): In some members VB are bicollateral. During
secondary growth internal phloem (primary phloem) is pushed near the centre of the
axis forming innermost part of the vascular tissue. This strand of phloem is termed as
intraxylary phloem or internal phloem. In most of the cases internal phloem is
primary. Common examples are- Argyreia, Calotropis, Leptadenia, Solaum, etc.
Interxylary phloem (Included Phloem): This type of secondary growth results in the
inclusion of strands of secondary phloem in the xylem. This may occur due to –i)
small segments of cambium producing phloem elements towards innerside for a short
period instead of forming xylem e.g. Combretum and Entada. and ii) small segment
of cambium cease to function. A new segment is formed just above the phloem and
function normally, thus embedding the phloem elements lying below, into the
secondary xylem. e.g. Strychnos. Included phloem occurs in Bougainvillea,
Leptadenia, Salvadora, etc.
11
Practical 3
PROPER USE OF A MICROSCOPE
1. Types of microscope
1. Simple microscope (Dissection microscope)
2. Ordinary light or compound or research microscope (Monocular / binocular /
trinocular)
3. Stereomicroscope
4. Polarization microscope
5. Phase contrast microscope
6. Fluorescence microscope
7. Electron microscope
SIMPLE MICROSCOPE
Understanding its construction / Parts
This type of microscope is use for the dissection, especially during anatomical
studies, embryo separation, etc. It consists of basal foot and a short limb. The stage is a
simple glass plate. A mirror is fixed below for the light adjustment. A folded arm is
attached to the limb which can be vertically moved by a rack and pinion system.
How to work
Eye is placed close to lens. The lens is moved over the glass plate and adjusted on
the object by tilting the folded part of the arm as desired. Object is now illuminated by
suitably turning the mirror below. An object is focused by the adjustment screw. In order
to get a clear view, lens and glass plate stage is always kept clean.
COMPOUND MICROSCOPE
Construction / Parts
i.
Foot and limb: Bottom most parts of
the microscope is the foot from
which arises the limbs
ii.
Stage: Stage is borne on limb and on
it objects to be viewed or examined
are placed. Two stage clips are also
provided to hold slides in position. In
research microscope there is always a
fixed mechanical stage. With the help
of a mechanical stage knob, slides
could be moved with much precision.
12
iii.
Mirror: Its location is below the stage, its function is to reflect light and illuminate the
objects
iv.
Body: Above the stage is the body. It is a brass tube which is movable by a rack and
pinion mechanism vertically. This is known as course adjustment. A fine adjustment
knob is also attached to a course adjustment knob / body and is used for finer
adjustment while using high powers.
v.
Lens system: are of two types
vi.
Eye piece or ocular: This is fixed on upper side.
Normal magnification is 10X but varied from 3X
to 15X .The oculars should be adjusted to suit
both of your eyes. Note that there is a scale on
the tube holding both objectives. We will label
microscopes so that each student can work with
the same instrument throughout the course.
Grasp the adjustable knurled ring below each
ocular with your thumb and forefinger and gently
rotate it so that each is set at 64 which is its midpoint. Before you make any
adjustments, place a slide on the stage and focus on part of the specimen.
vii.
Objectives & Nose Piece: Nose piece is meant for fixing and changing objectives.
Objectives are screwed at the lower end. The function of the objective is to form an
enlarged image of the object; whereas the eye
piece magnifies this image. The magnification
of an image is primarily controlled by the
objectives which are housed in a rotating nose
piece. To change objectives you rotate the
nosepiece, starting with the 4 X objective. Do
not start viewing by swinging in the 20 to 100
X objectives. These may be damaged if they hit
the specimen. Objectives of varying focal
length vis–a-vis magnification are available.
Low power objective (e.g. 4 X or 16 mm) have focal length of 2/3 inch while high
power objectives such as 10X or 4 mm, 40X and 100X have relatively longer focal
length (>1/6 inches). An oil immersion lens (100 X or 2 mm) is also fixed to the
objectives of 1/12 inch focal length. The magnification is indicated by a number on
each objective. Furthermore there is a progression in size such that the longest
objectives have greatest magnification. The distance between the objectives and the
cover slip (working distance) decreases dramatically as the magnification of the
objective increases. The 100 X objective is an oil immersion lens. Note the black line
near the tip of the objective. This is used to identify an oil immersion lens. Place a
small drop of thickened cedar wood oil on the objective lens. A small drop of oil
must also be placed on the cover slip. The lens should be carefully lowered into the
oil prior to focusing. In this case use only finer adjustment very slowly and carefully.
It is very useful for bacterial and cytological studies. Oil improves the optics because
it unites the glass cover slip and the objective. It replaces air with oil. The oil has the
same refractive index as glass. Thus less light scattering & refraction occurs. Be sure
that the specimen was in focus at 40X before switching to 100X.
13
viii.
Sub-stage condenser: It is used for concentration of light on objects. It has got an iris
diaphragm for controlling the light and a frame for
fixing filters.
HOW TO ADJUST IRIS AND DIAPHRAGM
The best resolution occurs when all elements of
the microscope are in perfect alignment and the iris
diaphragms are properly adjusted to the best aperture.
1] Because we will be using a lot of thick hand-sections in this class, it is vital that you
learn how to achieve best illumination. Otherwise, you will not be able to analyze your
specimens Place a commercially prepared slide on the stage.
2] Make sure the swinging lens is in the light path (facing up) and focus on the specimen
using the 10X objective.
3] Use only one eye [right eye with right ocular or left eye with left ocular] and focus the
specimen with the coarse/fine focusing knob.
4] Use the knurled ring below the other ocular to focus it while
looking through it with your other eye. You may not need to change
the focus. However, experiment by rotating the knurled focusing
ring to see its effect. My German friends have told me that the
correct way to focus the second ocular is to make it more negative
so it is out of focus, then rotate it in a positive direction until it is
focused.
5] Having the oculars focused will improve image quality and will
decrease eye strain. Once this is done it need not be changed during
a given session. However, it is a good habit to do this at the
beginning of each lab. It is best done at 10X because there is less
chance for errors at this magnification compared to 4 X.
6] Make sure the aperture iris is completely open [rotated all the
way counter-clockwise].
7] Reduce the field of illumination by rotating the knurled ring on
the field diaphragm completely clockwise. Be gentle with the field
diaphragm. It should close without any effort.
8] You should see a small circle of light. If you are lucky, it will be
in the center of the field. However, it will most likely be off-center
and out of focus. Let us know if you can'
t find it!!
9] Use the vertical condenser adjusting knob to make the circle as
small as possible by gently rotating it. This moves the condenser up
and down. Do this carefully so that the circle of light is not pushed
laterally. As you focus the field diaphragm you will notice that its
halo turns from blue to red and red to blue. The best focus occurs when you adjust the
14
condenser so that the halo is just between red and blue. This is a little hard to do so don'
t
be too worried if you have some red or blue in the halo.
10] Expand the field diaphragm by rotating its knurled ring counter-clockwise, until the
light touches one edge of the field. If the light is perfectly centered it should touch the
entire circumference of the field. This is unlikely.
11] Center the circle of light by using the two small adjustable knobs on the front of the
condenser. When you are satisfied, expand the field so that the light fills it completely.
However, do not fully open the field diaphragm. Open it just enough to extend beyond
the field of view.
12] Repeat this with the 20 or 40 X objective. For critical work this should be done for
each objective. This is especially important for taking photographs and for examining
minute, translucent specimens like fungi and algae. For our labs, it will be good to do
this for the 10X objective at the start of each session. You need not do this for 4X and
40X. However, if you are having some problems resolving details, check to be sure that
you have the condenser aligned and focused.
It may be difficult to do this with the 100 X objective. However, if you achieve proper
alignment with the 40 X objective, the 100 X will be similar.
13] When working at 20 - 100 X it is important to adjust the condenser aperture iris. This
is especially important for translucent structures. Closing this iris increases contrast.
Thus something fuzzy becomes smooth and something faint becomes dark. It is usually
possible to close the iris and judge its effects subjectively. However, there is a "tried &
true" procedure which you should know.
14] Remove one of the oculars and look directly down the tube at the light field. Close
the iris so that it occludes 1/4 - 1/3 of the area. This should give the best contrast.
Examine a specimen before and after adjusting the aperture iris. This should be done for
each objective for critical viewing. In practice, you can experiment with this while
viewing a specimen and adjust it without removing the ocular. Closing the aperture iris
also increases depth of focus up to a point. Thus, more areas of a three dimensional
specimen will be in focus If it is closed to much, a flat indistinct image results.
The example shows part of a diatom frustule. There is little detail when the iris is wide
open (top). When it is fully closed (middle) the contrast is increased but there are
aberrations which make the small holes appear larger than they are in actuality. The
outline of the small holes is also indistinct. When the iris is closed 25 - 30 % there is
improved contrast and less aberration.
15] Experiment with the aperture iris while viewing a prepared slide. Once you have
achieved what you think gives the best image quality, remove one of the oculars and see
how much of the field is occluded.
Mechanical operation / working
15
1. Work with concentration
2. Have a good light-a north light (take seat facing North) in day time is best. Limb
must be toward you. If light source is artificial filter (preferably blue) be used
3. Focus light with concave mirror till you get the best light. It can be adjusted further
by moving the sub-stage up and down as well as, as with the help of iris diaphragm.
Use the plane reflector mirror if there is a sub-stage condenser and then focus the
condenser in such a way that either the image of the lamb or some distant object in
day time appear very clearly in the field of view. This is the correct position for
condenser to give critical illumination. In lower power if you see the image of
objects then insert a piece of ground glass in between the mirror and light. But
position of condenser should not be disturbed.
4. A prepared slide is place on the stage. Object is adjusted just over the stage aperture
5. Always start observing the object at lower power. If higher magnification is
desired, turn the nose piece to next high power. Work carefully with high power
and do final focusing now with fine adjustment. View through high power after
placing cover slips over the object..
6. Keep both eyes open. Practice to observe objects with both eyes. But while using
camera lucida it is convenient to use the left eye.
7. Additional work: Dark ground illumination produces beautiful effects. A small
disc of black paper is inserted to the lower side of the sub-stage condenser. It
blocks the central rays of light and only allows the peripheral rays to fall on the
object as a result of which the object appears to be self luminous. Coloured light
can also be used. This method of illumination is used for observing bacteria and
protozoa.
Magnification
The sum of magnifying powers of both objective and eye piece constitute the total
magnifying power of both microscope. These can be varied by changing one or the other.
When you present a photograph, taken from a microscope, mention its magnification at the
lower end of image (as x10, x40, x100, etc).
STEREOSCOPIC MICROSCOPE
It consists essentially of two microscopes of the same magnification arranged side
by side in collimation. Optical axes of the two microscopes are at an angle to each other,
and the specimen is viewed simultaneously through them. The two eye piece serving
together as binocular. It has one fixed objective lens and one or two eye pieces the powers
of which is changeable. Standard magnification of eye piece is 10X. The stage in the stereo
microscope is non movable but nose can be moved up and down on the limb.
Stereomicroscope has the provision for the diopter adjustment and the interocular distance
can be set according to the suitability of the observer. The coincidence of the two images,
obtained independently by angular vision, produces an image that appears three
dimensional rather than flat causing a stereoscopic effect in the binocular view. This
16
microscope is used for observing three dimensional view of a small object and the object is
usually magnified up to 30 X.
Working in a stereo microscope
1.
Place the microscope where light reaches to its maximum or use the artificial
light source with blue filter or with fluorescent light
2.
Place the object (not minute anatomical section) either on the slides or directly
on the stage (do not make it dirty). If specimen placed on slide then keep the
slides on the stage.
3.
Move the nose up or down with the help of course adjustment knob while
viewing through eye piece.. Once specimens are visible or appear through the
eye piece then go for fine tuning with finer adjustment knob.
Precautions and Care
1. Never use direct sunlight as it is injurious to the eyes
2. Lens must be cleaned with fine quality tissue paper wetted with a drop of xylene.
Or with chamois leather or a soft silk handkerchief or mulmul or rubia violet piece.
Never touch the lenses with fingers
3. Fix the eye piece at their proper places. Never remove objectives from the
microscope. Keep microscope covered when not in use.
POLARIZATION MICROSCOPE
It is used to detect and analyse the birefringence of minerals or other materials.
(When an ordinary beam of light waves are vibrating in all planes along an axis passes
through a birefringent (anisotropic) object at an angle, it is split into two rays whose
vibrational planes are perpendicular to each other the phenomenon is called birefringence).
The microscope has two Polaroid filters: one below the condenser (polarizer filter) and the
other (analyzer filter) placed either in a slit in the mechanical tube between the objective
and the eye piece oe in the eye piece. The eye piece of an analytical polarization
microscope commonly has adjustable cross hairs. The stage is rotatable against a scale so
that the angular degrees of rotation can be determined. For the student’s practical class it is
not required hence the details are skipped.
Polarizing Filters cause light to vibrate in one plane. Light traveling along a
straight line vibrates in all possible planes. Imagine many radii emanating from a
common center. These would represent the many vibration planes of the light beam. A
polarizer cuts out all but one of these. If two polarizer are oriented at 90o to one another,
no light will pass through the second one in the series. Verify this by holding one
polarizer while looking at a bright object. Take a second polarizer in your other hand and
superimpose it on the first. Turn either one until the light is completely blocked. If a
17
crystalline or paracrystaile object is placed between crossed polarizer, it will depolarize
the light which passes through it. This property is known as birefringence. Consequently,
the birefringent material will be visible while all else will remain dark. Cell walls,
crystals and some starch grains are birefringent, and become apparent using polarized
light. This works with unstained and stained sections.
Amyloplasts from Cana seen with
typical bright-field illumination
Amyloplasts from Cana seen with crossed
polarizer
PHASE CONTRAST MICROSCOPE
FLUORESCENCE MICROSCOPE
Fluorescence microscopy is based on the property of certain substances to get
excited when irradiated with high energy radiations of shorter wave lengths (such as UV,
blue violet and blue) and emit low energy light of longer wavelengths (such as green,
yellow and red). The emission ceases within 10-9 seconds of the removal of the exciter
radiations and is referred to as the fluorescence. Fluorescence microscope has two filters: i)
exciter, and ii) absorption filters. exciter filters are placed in the optical axis of the
microscope between the light source and the object. They help in the selection of the
desired exciter radiations by filtering away the unwanted wavelengths.
PHASE CONTRAST MICROSCOPE
18
Practical 4
MEASUREMENTOF MICROSCOPICAL OBJECTS AND DRAWINGS
Micrometry
Micrometry deals with the size measurement of microscopic objects. Measurement
scales used for this purpose are known as micrometers. These are of two types- one is stage
or slide micrometer and the other is ocular micrometer which is to be fixed in the eye
piece. Ocular micrometer consists of a glass disc with a graduated scale engraved in its
centre. The stage micrometer look like a slide in the middle of which 1 mm long scale is
engraved with 100 equal divisions, hence one division in stage is equal to 0.01 mm or 10
(1 micron=1/1000 mm). In fact this is also mentioned on one side the stage micrometer as
1/0.01. Numerical preceding the oblique sign represents the total lengths of the scale in
mm and the one that follows the sign, distance between the shorterst intervals in mm.
To measure the object, the division in the ocular micrometer has to be standardized
for every objective-eye piece combination and for every microscope you are going to use.
For this standardization, the stepwise procedure is given below.
1. Remove one eye piece from a microscope in which you want work (5 x or 10 x)
and place the ocular micrometer in the collar sleeve meant for this purpose in the
eye piece tube between the top and bottom lense.
2. Place the stage micrometer (slide) on the stage
3. Adjust the focus by moving the stage or nose up and down till you get clear image
of marking on the slide micrometer
4. Now superimpose the two reading i.e. eye piece marking over the markings of
stage micrometer and find out the two exactly coinciding divisions on the ocular as
well as stage micrometer
5. Count the number of divisions between the two coinciding lines on both ocular and
stage
6. Now move the stage micrometer to a new position on the stage and repeat the step
3 to 5 and count the number of division between the two coinciding lines on both
ocular and stage. Repeat the step again so that you will get five to ten counts
7. Tabulate the values and work out average number of division between two
coinciding lines on both ocular and stage micrometer
Count
No
1
2
Number of division on the Number of division on the stage
ocular
micrometer micrometer between two coinciding
between two coinciding lines
lines
19
3
…
Mean
8. Find the value of one ocular division by using the formula i.e.1 ocular division =
mean number of division on stage divided by mean number of corresponding
division on ocular multiplied by 10 and express the value in micron. For example,
if 8 divisions of the stage micrometer correspond to 10 division of the ocular, then
one ocular division would be 8/10x10 = 8 micron
9. Once you know one division of a ocular in terms of micron, then remove the stage
micrometer and in that place keep your specimen and view through the ocular.
Measures the length or width of the object in terms of number of ocular division
and then multiply this values with calibration factor, you will get exact length or
width of your microscopical object.
Note:
The calibration you did is specific for a particular microscope and objective lens.
Therefore, when you desire to use all the objective lens in a microscope, do calibration for
different powers (magnification) separately. Once calibrated the same value can be used
repeatedly for that particular microscope and that particular power.
HOW TO DRAW A DIAGRAM
Drawings from microscopical observation are of three types
1. Diagramatic sketch: This means a schematic representation of different tissues by
lines, dots, crosses and other signs. Never draw cells in a diagrammatic sketch
2. Drawings or detailed sketch or cellular sketch: In these details of cell
construction is done as accurately as possible. Never be schematic or use signs in
cellular sketch and never make drawings very extensive. Rather select such a sector
which includes all important cells, tissues and structure. Cells at edges always must
be left unfinished to show continuity.
3. Camera Lucida drawings: This instrument is provided with a reflecting prism and
a mirror and is indispensable for any researcher when true to scale drawings are
needed. All your figures should be graphic records of your observations. They
should be drawn with pencil. They should not be copied out from text books.
Drawings and diagrams should be labeled properly
Note: Use Indian ink (black) and fine pointed special drawing pen for making images. It is
advisable to draw the image in special quality thin drawing papers
20
Practical 5
UNDERSTANDING PHOTOMICROGRAPHIC CAMERA SYSTEM
This unit is composed of a typical light microscope with a trinocular head that has
two oculars and a photo-tube.
There is a projection ocular in the photo-tube. This focuses the image onto the film.
Light is simultaneously transferred to the
oculars & the photo-tube.
The photo-tube has a 35 mm camera back
that is simply a film holder.
There is a separate light meter & shutter
control box. This is used to select the
correct film speed (sensitivity). The shutter
button is located here. Turn this unit on
using the switch on the back.
There is also a separate illuminator for the
microscope.
To insert a roll of film, turn the transfer
lever fully clockwise. It will not become
completely horizontal. Do not exert force on
it!
Pull out the light blocking slider.
Pull out the rewind knob & the camera back should open.
Carefully open this all the
way.
Place film in the right-hand
compartment (as you face the
camera).
Stretch the film so that the
emulsion side is down & it
completely covers the film
uptake sprocket.
one of the scale-like flaps on the film uptake sprocket.
Slide the end of the film under
Be sure to engage one of the holes along the film margin with the small hooks on the film
transport sprocket.
21
Press the manual film advance button & advance the film with the manual film transport
lever to firmly attach the film to the film
uptake sprocket.
Close the camera back & press the advance
button & advance the film twice. This will
remove partially exposed film.
Select the appropriate film speed on the light
meter shutter control box.
Use the film speed selector knob to rotate the
settings to match the film speed with the 35
mm reference mark on the right side of the knob.
The
American
units
are (ASA) 100, 200, 400, 800.
The European units (DIN) for these are
21, 24, 27, 30. USE ASA 100 (DIN 21)
color film gives high quality images.
The 35 mm film selected in the illustration
has a din of 12 and an ASA of 12.5.
Slide film gives the best resolution but
print film is more forgiving in terms of
acceptable exposures.
Set the shutter speed selector on the lower
right side of the shutter control box to
auto!
The photo reticle move the specimen out of
the way.
Look through the right ocular to observe
the photo reticle.
The light gray rectangle with dark corners, shows the area of the image with the 16 mm
projection lens.
The darker, inner rectangle encloses the focusing cross hair and does not refer to any image
size.
The outermost, incomplete rectangle is for the 10x projection lens which we do not have.
Note the focusing cross hair in the center of the photo reticle!
22
Focusing the ocular with the photo reticle. This is a most important operation! If this is not
done correctly, all of your photos may be out of focus. Defocus the right ocular by rotating
it fully counterclockwise. You will need to hold the lower, outer part of the ocular and
rotate the upper, inner cylinder that holds the ocular lens. Otherwise the whole thing will
rotate and go round and round and round.
Carefully rotate the ocular clockwise until
the cross hair at the center of the reticle can
be clearly resolved with your right eye. See
the example on the right.
Taking a picture
i. Place the specimen under the
objective to compose the picture.
ii. Adjust the condenser iris (10x and
above).
iii. Turn the illumination up so that the
needle in the meter coincides with
the red line. This will insure that the
correct "color temperature" is
achieved.
iv. Do not look through the oculars with
the light set to the red line!
v. Push the shutter release button on the light meter.
vi. The shutter should open and close quickly!
vii. Do not look through the oculars with the light set to the red line!
viii. Lower the illumination to spare the bulb and to safeguard your eyes.
ix. Click the manual film advance button and use the film transport lever to advance the
film prior to taking the next exposure. Otherwise you will get a double exposure!
Rewinding & removing film
i. When all of the film is exposed the
"end of roll light" should be
illuminated.
ii. Insert the light blocking slider all the
way.
iii. There is no rewind lever!
iv. Locate the silver rewind button on
the left rear of the camera body &
depress it!
v. Pull out the silver rewind handle
from the rewind knob. Do not pull
out the entire knob!
vi. Rotate the rewind handle clockwise until the tension is released.
vii. Keep rotating to be sure that all of the film has been rewound.
viii. Now Pull out the rewind knob.
ix. The camera back should pop open.
x. Open it all the way and remove the exposed film cassette.
23
Practical 6
PREPARATION OF GLASSWARE CLEANING SOLUTIONS
The following solutions can be used for washing glassware and slides.
1) Chromic acid preparation
Reagents:
Chromic acid (K2Cr2O7)
Water
Conc. H2SO4
= 63 G
= 35 ml
= 960 ml
Method: Potassium dichromate is dissolved in water taking in beaker and
transferred to glass trough. Sulphuric acid is poured slowly and stirred / shaken
intermittently.
Caution: Be careful while adding Sulphuric acid to water
2) Nitric acid and Hydrochloric acid mixture
Reagents:
Conc. HNO3
Conc. HCl
= 1 part
= 4 parts
Note: This mixture does not last long & unpleasant
3) Method of washing:
First pour the solution into a plastic tray or enamel coated tray and then place
the glassware for an hour or so. Do not use your hand immediately for washing. At
the end of soaking period, pour water into the tray or decant the acid solution from
the tray and then again add water then with your hands rub and wash the glass ware
carefully.
If readymade solution like CLEANSOL or detergent is available use it for
secondary cleaning after removing the stains adhered on glassware with acid
solutions. Cleansol may be diluted 10 times before use. Whenever detergent is used
for cleaning, thoroughly remove the detergents by repeated washing in tap water.
Note: Teacher may instruct the laboratory assistant to follow the above method for
washing and with precautions
24
Practical 7
PREPARATION OF MATERIALS FOR ANATOMICAL STUDIES
The materials to be preserved should usually be fresh. The three different processes
which are involved in the preparation of materials may be done either in one combined
operation or separately.
1. Fixing: It consist of i) very rapid killing of the protoplasm of the cells or
terminating the life processes in the tissue and ii) halting the postmortem changes
or the putrefaction, liable to occur in a dead system.
2. Hardening: This is required for soft materials to make the cutting of section
possible
3. Preserving: Preservatives are used so that materials are not degenerated
FIXATIVES
A fixative is a reagent which fixes or stabilizes the living tissues. A variety of
fixatives are available. Each one has some merit at the same time there is no fixative is
perfect and universal. The best fixative is one which halts the life process without
structural disturbance or minimum distortion of the arrangement of the cells in the tissues.
The followings are recommended for anatomical studies.
1. Formalin Alcohol (Chicago Formula or Acetic Alcohol or Farmer’s fluid)
Absolute Alcohol
Formalin
= 100 ml
= 6 ml
Materials may be fixed for 10 hours and stored in this fluid indefinitely. Fixation in
this is recommended for the localization of insoluble polysaccharides as it dose not contain
water hence retain sugar but do not use this if protein is to be localised.
2. Carnoy’s fluid
Two versions are available; second version is popular which contains
Alcohol (100%)
Acetic Acid (Glacial)
Chloroform
= 30 ml
= 5 ml
= 15 ml
Duration of fixation: 15 min – 24 hrs. Store the materials in 70 % alcohol after
washing. It is preferred for studies on chromosome and nucleic acid. Good for squash and
block preparation of root tips, anther and buds. It has great penetrating power due to
inclusion of chloroform.
25
3. Formalin- Aceto-Alcohol or FAA
Alcohol (50-70%)
Acetic Acid (Glacial)
Formalin (40 %)
% formaldehyde)
= 90 ml
= 5 ml
= 5 ml (Formalin is the trade name of 38-45
It is a Universal fixative or standard preservative used in histological studies and in
some histochemical studies like localization of proteins and polysaccharide. For woody
materials such as stem which needs rapid penetration the proportion of Formalin may be
increased and Acetic acid decreased. Materials may be fixed for 18-48 hrs.
4. Formalin- Propiono-Alcohol
Similar to FAA but use propionic acid instead of acetic acid
5. NKL fixatives
Acetic Acid (Glacial)
Formalin
Chromic acid
Water
= 10 ml
= 40 ml
=1g
= 100 ml
6. Zinc Formaldehyde
ZnSo4
Formalin (40 %)
Water
= 25 g
= 10 ml
= 100 ml
7. Randolph’s modified Navashin fluid
Solution A
Chromic acid
Glacial acetic acid
Water
=5g
= 50 ml
= 320 ml
Solution B
Natural formalin
Saponin
Water
= 200 ml
=3g
= 175 ml
Mix equal amount of Solution A and B when ready to use. buds, root tips and
similar objects can be fixed in this fluid.
8. Sorensen's Phosphate Buffer
Stock solutions:
Stock A: 0.2 M solution of NaH2PO4.H2O (27.6 g/liter)
Stock B: 0.2 M solution of Na2HPO4
26
To make 0.2 M Buffer for mixing with fixatives: mix 23 ml stock A and 77 ml
stock B. ph 7.3.
To make 0.1 M Buffer with for washes: mix 23 ml stock A and 77 ml stock B. Add
100 ml deionized H2O. pH 7.2 - 7.3.
9. Neutral buffered Formaldehyde
Usually 4 % Sol. of Formaldehyde in water or in phosphate buffer at neutral pH is
used for fixation of materials meant for localisation of protein and other histological
studies. In some studies it is not recommended to use commercial Formalin. Therefore,
prepare the solution as directed below and use afresh. The procedure for preparing 50 ml of
4 % Formaldehyde in 0.1 M phosphate buffer at pH 7.2 is given below.
Solution A: is prepared by dissolving 2 g paraformaldehyde powder in 20 ml of water by
gently heating (80 oC) and stirring with a glass rod. Add drop wise a 4 % solution of
Sodium hydroxide until the solution becomes transparent, making a total volume of 25 ml.
Solution B: Dissolve 0.636 g disodium hydrogen orthophosphate and 0.194 g potassium
dihydrogen orthophosphate in 25 ml water.
Mix equal volume of A and B and adjust pH to 7.2.
General procedure for fixing plant materials
i. Collect the live specimen from the field (e.g. root, shoot, bud, fruit, etc) and wash
to free derbies and soil particles adhering on it
ii. If the specimen is large cut into small pieces
iii. Immerse the specimen / tissues in a suitable fixative at room temperature for the
time specified under each fixative or for 10 hrs to overnight. For small objects not
exceeding 1 mm3 in size, fixation time as short as 2 to 4 hrs may suffice
iv. At the end, remove the tissues from fixative; do not through the fixative; can be
reused. Use small size filters or forceps to take out the tissues from fixatives. Use
hand gloves to avoid the fixatives spilling on your hands
v. Wash the tissues first in water and then 3 to 5 changes in 70 to 90 % ethanol.
vi. Storage of the fixed tissues, if desired, may be done in 1 % v/v glycerol prepared in
70 % aqueous ethanol or
vii. Proceed with infiltration and embedding or smearing
Caution: All of these except the Phosphates are hazardous and require special handling.
Note:
Instant killing of the material is most preferred. Therefore carry the fixatives to the field.
•
•
Acetic acid causes swelling and partial dissolution of the cytoplasm.
Formalin is slow and does not harden.
In general, the following precautions must be observed in fixing, hardening and
preserving the materials.
27
•
•
•
•
Firstly materials should be fresh
secondly, pieces must be cut as small as possible
thirdly a large bulk of the fluid must be used and
Completely immerse the plant material in the killing agent.
Creation of vacuum for microtome sectioning
•
•
•
•
•
•
•
It is desirable to draw a vacuum on freshly fixed samples. This helps the
fixative penetrate the specimen completely.
There are small, plastic hand vacuum pumps and desiccators that are light
and easy to transport.
Leave the container lids loose during evacuation.
OR place specimens in plastic scintillation vials & seal them with Parafilm.
These vials are cheap and rugged and you can reuse the packing materials
that are used for shipping.
Larger bottles may be required.
Avoid GLASS containers or other easily broken material.
Test them to be sure they won'
t leak or dissolve.
HARDENING
For most botanical materials strong alcohol 70 % or upward does all the hardening
required; 70 % is perhaps the best all round reagent. For delicate tissues very strong grades
are harmful. Complete dehydration is necessary before embedding in paraffin wax for
microtomy.
DEHYDRATION BEFORE MOUNTING & PRESERVATION
1. After fixing and washing, the materials must be brought upon strong alcohol by
stages. Starting from water the materials should pass through grades of alcohol such
as 10 %, 30 %, 50 %, 70 % and 90 % in successions and allow the materials about 2
hrs in each strength. Preserve the materials in 70 % alcohol.
2. Instead of Alcohol, Xylol series is also recommended for dehydration
3. Remember to always cover the trough containing alcohol series while the specimens
are in or out
28
Practical 8
UNDERSTANDING SOME ANATOMICAL TECHNIQUES
(Maceration, Peeling and Smearing)
1. MACERATION
The organs of plants contain many types of tissues. These cannot be seen by
sectioning or clearing. The three dimensional and real natures of the cells composing an
organ is understood by a special method known as maceration. This consists of isolating
individual cells from a mass of cells. In this technique, the middle lamella of the cell walls
is dissolved thereby allowing the cells to fall apart. The classic fluid for this purpose is
Schultze’s solution. Ammonium oxalate method is also preferred. Harlow’s method is also
in vogue. Alternatively, soak the materials in 5 % chromic acid overnight and then tease
out on slides.
Jeffrey’s method
1. Cut the materials, fresh or dried, into small slices thinner than a toothpick.
2. Boil the materials in water till it settle down at the bottom indicating that it is free
from air.
3. The materials is now placed in a test tube containing macerating solution (contains
equal volume of 10 % Nitric acid and 10 % Chromic acid)
4. Heat this fluid and watch the condition of the materials by piercing it with the
needle. Stop heating as soon as the materials become soft and pulpy.
5. Transfer the fluids to a watch glass and wash the materials repeatedly with water
till all the traces of acids are removed
6. The materials are now stained with safranin, de-stained and mounted in glycerine
or glycerine jelly.
7. If desired materials may be dehydrated, by passing through the alcohol series for
dehydration, followed by a graded series of xylole for clearing, before mounting in
Canada Balsam
Schultze’s method
1. Materials is kept for a few days in a macerating solution consisting of conc. HNO3
50 ml and 1 g Potassium chlorate (5 %) crystals or materials is sliced and boiled in
the solution for a few minutes i.e. till the materials is bleached white.
2. Remove or decant the acid from the materials; washed thoroughly till the materials
is free from all acid traces.
3. Tease or crush the materials till individual cells appear isolated and
4. Mount in glycerine
Harlow’s method
1. Sliced and boiled materials are treated with chlorine water for 2 hrs.
2. Wash with water
3. Boil the materials in Sodium sulphite for about 15 min
29
4. Drain out the liquid from materials and wash repeatedly with water
5. Teas or crush with needle or glass rods
6. Prepare either temporary mount in glycerine or permanent mounts in glycerine after
passing through alcohol or Xylol series
Ammonium oxalate solution
Hydrochloric acid
Alcohol (95 %)
= 30 ml
= 100 ml
1. Materials are kept in the above solution overnight.
2. It is then thoroughly washed and is kept in a 0.5 % Ammonium oxalate solution
and boiled for few seconds.
3. Mount in the same liquid i.e. 0.5 % Ammonium oxalate
2. PEELING
In order to study the number, arrangement, distribution and structure of stomata, leaf
epidermis is stripped off. The method consist of,
1. Break the leaf irregularly with a force. This easily separates a little part of the lower
epidermis which remains protruding on the lower surface of leaf
2. Pull the broken membranous piece so that a long ribbon of lower epidermis gets
removed
3. If lower epidermis does not easily separate, a needle or forceps is inserted and a
small part is slowly separated. Hold this piece in hand and pull apart a large strip.
4. Stain the lower epidermis (strip) with safranin and washed mount in glycerine or
glycerine jelly
5. If permanent preparation is desired, normal procedure of dehydration and clearing
is followed before mounting it in Canada Balsam
3. SMEARING
The principle underlying this method consists of spreading out the cells in a single
layer. Almost all the cells remain adhered to the slide. The cells are smeared at a stage
when they are in the process of cell division. This permits the study of various stage of cell
division and structure of chromosomes. Pre-requisite for such studies is the killing of
dividing tissue at the proper stage of cell division and selection of material where cells are
not firmly united with one another by middle lamellae.
30
Microsporocytes of Trillium, Lilium, Oenothera species as well as anthers of
Tradescantia, Triticum and Nicotiana species and root tips of onion, Ficus, etc. fixed at
opportune time are widely used for smear preparations.
Procedure
A procedure for taking smears from anther is described below.
This is a non-
standardized procedure
1. Slides should be perfectly clean for preparation of smears. For that immerse the
slides in glassware cleaning solution an hour ahead of your practical. Then clean
the slides with running water and clean and soft cloth or low cost tissue paper
2. Fresh anthers from buds are placed in the centre of slide.
3. The material may be killed in a fixatives.
4. To proceeds for killing, crush the anthers with scalpel or just using another slide
5. The slide is now inverted over a Petri dish containing killing fluid (fixative), in a
way that smeared surface comes in contact with the fluids.
6. Allow the slide in this position for 10-15 minutes; slide is now inverted with
smeared surface upward
7. Stain the slides
8. Sometimes after crushing the anther it is immediately stained without killing.
31
Practical 9
HOW TO MAKE A HAND SECTION
Anatomical technique like smearing cant be used when the cells have to examined
in spatial relationship with each other in the tissues, Further large organs or tissues
requires to be dissected into very thin slices in order to expose their inner details and to
increase their refractive visibility for the microscopic observation. Section cutting or
sectioning is the most common technique of studying microscopic anatomy or histology of
large specimens. Sectioning can be done using microtome or by hand. Free hand section is
the easiest, cheapest and fastest way of cutting specimen and for this the procedure is
given.
Instruction for Right-Handed sectioning
1.
Before you start your exercise, place a band-Aid on the thumb of your left hand.
Have the cotton portion on the bottom of your thumb. The thumb is a backstop for
this operation. Place another on the end of your index finger. The index finger will
control the height go the specimen, and thus its thickness
2.
Grasp the plant structure between your left hand thumb and forefinger so that the top
of the specimen extends above the level of your forefinger
3.
Use pith to embed the material if it is thin, small and soft. Pith of potato tuber,
radish, carrot, papaya fruit, tapioca, Pennisetum, etc are commonly used. Trim a
carrot to a size that is easy to hold and is large enough to hold the specimen. Blot the
outer cut surfaces so that they are not slippery. Make a slit or opening in the carrot
that will accommodate the specimen. Place the specimen into the slit. Squeeze the
carrot with your fingers to secure the specimen.
4.
For a beginner, razor is better than
safety razor blade since the former
is hard and inflexible and it will not
bend its course through hard
specimen. Razor is provided with a
handle and therefore has greater
maneuverability over the blades.
5.
If hone and leather strops are used,
hold the blade in such a way that the
blade and handle form right angle
with one another. The handle should
remain free, while index finger is
put on the hooked end of the razor.
First, second and third fingers are
pressed against the thick back edge
of the razor, while thumb remains pressed against the milled surface of thick shank
of the blade. f blades are used then hold a single edged blade in the right hand and at
right angle to the specimen so that uniform and thin sections are cut. Be sure that
blade is wet
32
6.
Raise the specimen slightly by manipulating it with your fingers and repeat the
slicing motion
7.
Thin sections can often be obtained by pressing the blade down on your forefinger
and then slicing through the specimen several times. The razor is then moved quickly
over the materials and a stroke is completed in one action.
8.
After several sections have accumulated on the blade, wash them off in a Petri dish
of water
9.
During this process of section cutting, both materials and razor should always be
kept flooded with water. Once sectioning is over, razor should be dried without
disturbing the edge, stropped, greased and encased.
10. If sections are thin, then float on the surface of eater. These are now selected and
placed on a clean slide and then observe under microscope. If this is a good and thin
section then use it for staining.
11. It is a good idea to view unstained sections prior to staining. Proper use of the
aperture iris is important for this.
Some tips
Considerable skill and experience are required for cutting satisfactorily thin and
even section. Don’t be frustrated with with your failure. Try and keep on try. You will get
good section nearing 25-30 thick, less than this thickness is impossible through free hand
section. Use Teflon-coated razor blades and platinum coated razors. The best blades are
those designed for old-fashioned double-edged shavers. These are very very sharp & must
be handled with great care. If you use hone and leather razor purchase hone from
Washitaw or Arkansas brand hones because their stone is fine grained)
Precaution: You might apply tape to one of the edges to avoid cutting yourself. Teflon
sprays can be purchased so that blades can be coated immediately prior to use
TIPS TO PLACE A COVERSLIP OVER A SPECIMEN
It is essential that the sections be completely immersed in water so that air is
excluded. Air bubbles or spaces will interfere greatly with your observations. To avoid
these when adding a coverslip do the following.
a] Place your sections in 2-3 drops of water or stain in the center of the slide.
b] Use a fine forceps to pick up a large cover slip (20 x 40 or 20 x 50)
c] Place one end of the coverslip on the slide (near boundary with frosting) without
touching the solution containing the specimens.
d] Steady this end with the fingers of your left hand.
33
e] Slowly lower the forceps until it touches the slide. By this time the coverslip should
have touched the solution on the slide.
f] Slowly remove the forceps so that the coverslip is gently lowered into its final resting
position.
g] Remove excess solution by touching the side of a paper towel to the narrow edges of the
coverslip. Be careful not to drag out your sections with the excess solution.
h] If you have been using a stain, add water to one end of the coverslip while withdrawing
the stain at the opposite end with a towel. In most cases you do not need to get all of the
stain out.
i] Wipe excess fluid from the bottom of the slide or it will stick on the stage and make your
life more miserable than it already is.
34
Practical 10
MOUNTING AND RINGING
The cut section or any preparation can be mounted temporarily or permanently.
For that different mounting media are available. They are Canada balsam, Glycerine,
Glycerine Jelly and DPX Mountant
1.
Glycerine: Pure Glycerine diluted to 15 to 25 % is widely used for mounting. Semi
permanent and temporary preparations are mounted in Glycerine only
2.
Glycerine Jelly: Gelatin – 1 parts; Glycerine- 7 parts; Water -6 parts. Warm the
gelatin for two hours by adding water and warm once again. Phenol 1 % is added
later. Crystals of Safranin may also be mixed if desired. Allow the solution to cool
and settle into jelly. It can be used for mounting any preparation without undergoing
the process of dehydration.
3.
Canada balsam: It is resin exudates from a conifer-Abies balsamea. Most suitable
for permanent slide preparation. Object can be stored in the same condition for as
many as 25 years at least. The materials to be mounted should come through Xylol
series. It has an optical property which allows perfect and clear view of an object.
4.
DPX Mountant: Commonly use for mounting permanent slides
HOW TO MOUNT
Temporary preservation of slides
Slides can be saved for short periods by sealing the edges of the coverslip with
freezing support medium or nail polish. The former used to stabilize tissues for cryosectioning and works well for saving slides. However, it only lasts for a day or two. If you
use nail polish, apply one generous coat and allow to dry. Then add a second coat. Staining
intensity will degrade over time.
Objects mounted on glycerine is a common practice in biological lab for short time
preservation.
Semi- Permanent mounting by RINGING
Temporary preparations can be made semi- permanent by sealing them with agents
like Canada balsam, gum dammar, nail polish, gold size, etc and is called ringing. Canada
balsam is most suitable among them..
1. Sealing is done by using ringing table or turn table. For this purpose round coverslip
are needed.
2. Ringing table consists of a metal disc which is movable and adjacent solid
immovable platform.
35
3. A fully mounted preparation is first adjusted over the metal disc. Position of the
cover glass be adjusted over the metal disc and slide is fixed in this position with the
clips provided on the disc for the is purpose
4. The metal disc is given a momentum with the left hand fingers. The right hand is
firmly placed on the resting platform of the table. This hand holds a brush dipped in
sealing medium. The circle of the coverslip is visible while disc is moving.
5. Touch the brush gently to the circle. There should be equal layers of sealing medium
around and above the coverslip.
6. Allow the slides to dry off the sealing agent
7. Once dry, another coat of ring may be applied. Repeat this step for 3 to times
8. When slides are dried scrap off the sealing agent spread over the slides by a sharp
blade.
9. If coverslip are square shape slides can be made semi-permanent by painting over the
edges with sealing agents.
Permanent mounting
1. Carry out Dehydration before Mounting. Pass the slide through a graded series of
Alcohol or Xylol i.e. 30 %, 50 %, 70 or 90 %. Remember to cover always alcohol
containing jars with lids.
2. Use clean preparation free from dust and finger prints. Hold on the edges of slides and
coverslip to avoid finger-prints on it.
3. Place the section / object in the centre of the slides
4. Add one or two drops of mountant over the specimen. Do not allow the mountant to
flow out of coverslip, spreading over the slides. Remove excess mountant by touching
it with apiece of blotter paper
5. No air bubbles should enter the medium while mounting. This result in drying of
medium and preparation becomes unworthy for observation.
6. To avoid air bubble, touch one side of the coverslip to the drop of mounting medium
on the slide. Coverslip is held with needle. It may now be lowered gradually and
needle is removed. This ensure clean preparation and can be perfected after a little
practice.
7. Preparations when complete label it on one side of a slide with date.
36
Practical 11
STUDY ON MICROTOMES
Introduction
In order to avoid inconsistence in section thickness and to enhance the
reproducibility of results, instrument were devised and were known as cutting engine until
1839 when Chevalier used the word microtome for them. Modern microtomes are
precision instruments designed to cut uniformly thin sections of a variety of materials for
detailed microscopic examination. For light microscopy, where magnifications can reach
up to 1,800 X, the thickness of a section can vary between 1 and 10 microns (thin
sections). For electron microscopy, where magnifications of several hundred thousands are
possible, the thickness of a section is usually of the order of 10 nanometres (ultra-thin
sections). Both thin and ultra thin section can be made through microtome.
Parts of a Microtome
All microtomes consist of three main parts:
•
•
•
Base (microtome body)
Knife attachment and knife
Material or tissue holder
Working principle
With most microtomes a section is cut by advancing the material holder towards
the knife whilst the knife is held rigidly in place. The cutting action which can be either in
a vertical or horizontal plane is coupled with the advance mechanism so that the material
holder is moved after each cut. The distance
moved is pre-selected using a scale setting
on the microtome body and usually extends
between 0.5 and 50 microns on microtomes
cutting thin sections and from less than 60
nm to over 500 nm on machines cutting
ultra thin sections.
Types of microtome and their uses
1. Rotary Microtome
This is a general purpose microtome
for cutting semi-thin to thin sections for light microscopy. The microtome operation is
based upon the rotary action of a hand wheel activating the advancement of a block
towards a rigidly held knife. The block moves up and down in a vertical plane in relation to
the knife and therefore cuts flat sections. Available machines range from lightweight,
rotary microtomes suitable for cutting paraffin wax embedded material (also resin
embedding) in a continuous ribbon to heavy duty, motor driven instruments used with a
slow, continuous speed and retracting advance movement to section hard material
37
embedded in synthetic resin. The rotary microtome can also be found in most cryostats for
cutting frozen sections. Section thickness settings range from 0.5µm to 60µm on most
machines. Sections of paraffin wax embedded tissues are normally cut within the range 3
to 5µm whilst resin sections are between 0.5 to 1µm. Rotary microtomes are especially
suited to cutting sections using disposable steel knives.
2. Sledge Microtome
These are designed for cutting large blocks of paraffin and resin embedded material
including whole organs, for light microscopy. The knife holding clamps allow the knife to
be offset to the direction of cut, a major advantage when sectioning large, hard blocks. The
microtome, which is very heavy for stability and not usually subject to vibration, can also
be used to cut materials from various industrial applications (wood, plastics, textile fibres).
They are not suitable for cutting very hard resins such as araldite because of the risk of
vibration.
The Sliding microtome uses a slicing motion to make sections while the rotary
microtome uses a chopping action to cut specimens. Slicing is preferable because it places
less pressure on the specimen at the point of contact compared to chopping. That is why
slicing is better than chopping for hand sections. The Sliding microtome is commonly used
for sectioning wood.
3. Freezing Microtome
This form of microtome is used for cutting thin to semi-thin sections of fresh,
frozen tissue and semi-thin sections from industrial products such as some textiles, paper,
leather, soft plastics, rubber, powders, pastes and food products. The freezing microtome is
equipped with a stage upon which tissue can be quickly frozen using either liquid carbon
dioxide, from a cylinder, or a low temperature recirculating coolant. Some cooling systems
also allow the knife to be cooled at the same time. The cutting action of the freezing
microtome differs from those described previously as in this case the knife is moved whilst
the tissue block remains static. The block moves by a pre-set amount, in microns, at the
end of each cut. Consistent, high quality, thin sections are very difficult to obtain with this
type of microtome.
4. Ultramicrotome
The ultra microtome is used to prepare ultra thin sections for light and electron
microscopy. Very small samples of tissue or industrial product are usually embedded in
hard resin before cutting. It has been reported that sections can be cut as thin as 10
nanometres. Two forms of advance mechanism have been developed in this style of
microtome. The thermal mechanism relies upon heat induced expansion in a bifurcated
metal strip whereas in the mechanical form a microprocessor coupled to a precise stepping
motor controls the advance mechanism. The cutting stroke is motor driven to provide a
regular, smooth motion for sections of even thickness and constant reproducibility. Knives
are usually made from glass, diamond or sapphire. The block is brought to the knife edge
under microscopical control and as each section is cut it is floated on to a water bath
adjacent to the knife edge.
38
5. Cryostat
The processing steps for normal microtome such as fixation, dehydration,
embedment, etc. are time consuming and often alter the cell structure in subtle ways.
Fixing cells with formaldehyde, for example, will preserve the general organelle structure
of the cell, but may destroy enzymes and antigens which are located in the cell. Valuable
time can be saved by skipping the fixation and dehydration steps required for paraffin
embedding, and freezing the tissue in a modified microtome, the cryostat. Additionally,
frozen sections will more often retain their enzyme and antigen functions.
A cryostat is primarily used for cutting
sections of frozen tissue as well as pastes, powders
and some food substances. The cryostat commonly
consists of a microtome contained within a
refrigerated chamber, the temperature of which can
be maintained at a preset level. A recent innovation
has the body of the microtome positioned outside
the refrigerated chamber. The cryostat usually
contains a rotary microtome although some
portable units utilise a rocking microtome. With the
object, object holder and knife all at the same temperature and all other conditions for
cutting the material optimal, sections as thin as 1 micron are possible.
6. Hand microtome
This has a central cylinder in which the specimen is
placed. A carrot or other material can be used to support the
specimen if it is necessary. The specimen is secured with a
Clamping Screw on the side of the microtome. Section
thickness is controlled by a knob. The scale reads in microns.
Sections are made by slicing with a microtome knife, straight razor or other suitable
instrument. The successful use of a hand microtome is limited to sectioning intrinsically
rigid botanical material. It is difficult to obtain thin, even sections of animal tissues.
39
Practical 12
PRE-TREATMENT FOR MICROTOME SECTIONING
FIXATION
Since cellular decomposition begins immediately after the death of an organism,
fixing is done to prevent alterations in the cell structure through decomposition. Routine
fixation involves the chemical cross-linking of proteins (to prevent enzyme action and
digestion) and the removal of water to further denature the proteins of the cell. Heavy
metals may also be used for their denaturing effect.
Small pieces of specimen are removed from plants and placed in the fixative. They
are allowed to remain in the fixative for a minimum of four hours but usually overnight.
The longer the blocks remain in the fixative, the deeper the fixative penetrates into the
block and the more protein cross--linking occurs. The fixative is therefore termed
progressive. Blocks may remain in this fixative indefinitely, although the tissues will
become increasingly brittle with long exposures and will be more difficult to section.
While it is not recommended, sections have been cut from blocks left for years in formalin.
Formalin has lately been implicated as a causative agent for strong allergy reactions
(contact dermatitis with prolonged exposure) and may be a carcinogen. It should be used
with care and always in a well ventilated environment. Formalin is a 39 % solution of
formaldehyde gas. The fixative is generally used as 10 % formalin or the equivalent 4 %
formaldehyde solution. The key operative term here is gas.
Caution: Formaldehyde should be handled in a hood, if possible. As a gas, it is quite
capable of fixing nasal passages, lungs and corneas.
Dehydration
Fixatives, such as formaldehyde, have the potential to further react with any
staining procedure which may be used later in the process. Consequently, any remaining
fixative is washed out by placing the blocks in running water overnight or by successive
changes of water and /or a buffer. There are myriad means of washing the tissues (using
temperature, pH and osmotically controlled buffers), but usually simple washing in tap
water is sufficient.
If the tissues are to be embedded in paraffin or plastic, all traces of water must be
removed: water and paraffin are immiscible. The removal of water is dehydration. The
dehydration process is accomplished by passing the tissue through a series of increasing
alcohol concentrations. The blocks of tissue are transferred sequentially to 30%, 50%,
70%, 80%, 90%, 95%, and 100% alcohols for about two hours each. The blocks are then
placed in a second 100 % ethanol solution to ensure that all water is removed. Note that
ethanol is hydroscopic and absorbs water vapor from the air. Absolute ethanol is only
absolute if steps are taken to ensure that no water has been absorbed.
40
It is important to distinguish between dehydration and drying. Tissues should
NEVER be allowed to air dry. Dehydration involves slow substitution of the water in the
tissue with an organic solvent. For comparative purposes, consider the grape. A properly
dehydrated grape would still look like a grape. A dried grape is a raisin. It is virtually
impossible to make a raisin look like a grape again, and it is equally impossible to make a
cell look normal after you allow it to dry.
Embedding
After dehydration, the tissues can be embedded in paraffin, nitrocellulose or
various formulations of plastics. Paraffin is the least expensive and therefore the most
commonly used material. More recently, plastics have come into increased use, primarily
because they allow thinner sections (about 1.5 microns compared to 5-7 microns for
paraffin).
The following is a guide to the thickness at which sections can be obtained from
different embedding media ranging from soft (gelatin) to hard (resin):
Gelatin - 50 to 200 µm
Ice - 5 to 20 µm (frozen section)
Paraffin wax - 1 to 15 µm
Paraffin wax/resin mixtures - 0.5 to 2 µm
Resin - 0.05 to 1 µm
Paraffin embedding is good if you have a lot of sectioning to do & if serial sections
are important. However, the paraffin method has given way to protocols that use resins.
Resin-embedded materials can produce thinner sections compared to paraffin. The paraffin
method destroys most cytological features and it causes swelling and shrinkage. Resin
samples contain much more cytological details. Paraffin embedding is also extremely
laborious and requires a plethora of ovens, organic solvents and other specialized
equipment.
1. Paraffin embedding
For paraffin embedding, first clear the tissues. Clearing refers to the use of an
intermediate fluid that is miscible with ethanol and paraffin, since these two compounds
are immiscible. Benzene, chloroform, toluene or xylol are the most commonly used
clearing agents, although some histologists prefer mixtures of various oils (Cedarwood oil,
methyl salicylate, creosote, clove oil, amyl acetate or Cello solve). Dioxane is frequently
used and has the advantage of short preparation times. It has the distinct disadvantage of
inducing liver and kidney damage to the user and should only be used with adequate
ventilation and protection.
Caution: Be wary of all organic solvents. Most are implicated as carcinogenic agents.
Heed all precautions for the proper use of these compounds.
The most often used clearing agent is toluene. It is used by moving the blocks into
a 50:50 mixture of absolute ethanol: toluene for two hours. The blocks are then placed into
pure toluene and then into a mixture of toluene and paraffin (also 50:50). They are then
placed in an oven at 56 - 58° C (the melting temperature of paraffin). The blocks are
41
transferred to pure paraffin in the oven for 1 hour and then into a second pot of melted
paraffin for an additional 2--3 hours. During this time the tissue block is completely
infiltrated with melted paraffin. Subsequent to infiltration, the tissue is placed into an
embedding mold and melted paraffin is poured into the mold to form a block. The blocks
are allowed to cool and are then ready for sectioning.
Plastic
More recent developments in the formulation of plastic resins have begun to alter
the way sections are embedded. For electron microscopy that requires ultra thin sections,
paraffin is simply not suitable. Paraffin and nitrocellulose are too soft to yield thin enough
sections. Instead, special formulations of hard plastics are used, and the basic process is
similar to that for paraffin. The alterations involve placing a dehydrated tissue sample of
about 1 mm into a liquid plastic which is then polymerized to form a hard block. The
plastic block is trimmed and sectioned with an ultra microtome to obtain sections of a few
hundred Angstroms. Table 1 presents a comparison of paraffin embedding with the typical
Epon embedment for TEM.
Table 1 Light and electron microscopy preparations.
Sample Size
Fixative
Post-Fixation
Dehydration
Clearing Agent
Embedding Material
Section Thickness
Stains
Light
1 cm
Formaldehyde
None
Graded Alcohol
Xylol/Toluene
Paraffin
5-10µ
Colored dyes
Electron
1 mm
Glutaradehyde
Osmium Tetroxide
Alcohol or Acetone
Propylene Oxide
Various Plastics
60-90 nm
Heavy Metals
Softer plastics are also being used for routine light microscopy. The average
thickness of a paraffin-sectioned tissue is between 7 and 10 microns. Often this will consist
of two cell layers and, consequently lack definition for cytoplasmic structures. With a
plastic such as Polysciences JB--4 it is possible to section tissues in the 1--3 micron range
with increased sharpness. This is particularly helpful if photomicrographs are to be taken.
With the decrease in section thickness, however, comes a loss of contrast, and thin sections
(1 micron) usually require the use of a phase contrast microscope as well as special
staining procedures. The sharp image makes the effort worthwhile. These soft plastics can
be sectioned with a standard steel microtome blade and do not require glass or diamond
knives, as with the harder plastics used for EM work.
Write TBA series and Xylen:Alcohol series
42
Practical 13
TAKING SECTION IN MICROTOME
SECTIONING IN ROTARY MICROTOME
Rotary microtome consists of a stationary knife holder/blade and a specimen holder
which advances by pre-set intervals with each rotation
of the flywheel mounted on the right hand side. A
control knob adjusts internal cams which advance the
paraffin block with each stroke. It is relatively easy to
section paraffin at 10 microns but requires a lot of
skill and practice to cut at 5 microns. Since each
section comes off of the block serially, it is possible
to align all of the sections on a microscope slide and
produce a serial section from one end of a tissue to
the other.
While virtually anyone can cut a section within minutes of being introduced to the
microtome, proper use of the microtome is an art form and requires practice and
inventiveness. The essential steps are,
•
Specimens are clamped in the specimen holder and the three clamping screws are
used to make the specimen parallel to the knife.
•
The knife is secured in the knife holder which has a knife angle adjuster on one
side.
•
Always carry a microtome knife by its screw-on Handle
•
The crank handle is rotated clockwise to advance the specimen.
•
Section thickness (microns) appears in a dial on the front of the microtome. There
is an external knob at the rear which changes the thickness setting. Start around 20
microns. The section will advance each time the crank is rotated
WARNING - Microtome Knives are EXTREMELY SHARP & Dangerous. They are so
sharp that you can hardly feel them cutting into your hand. Be constantly aware of
the blade'
s location! Always use a handle to carry a microtome knife. If the
microtome knife has contact with water, it needs to be dried and oiled ASAP
SECTIONING IN SLIDING MICROTOME
The Sliding Microtome has three main parts. These are, a] Knife holder b] Slide
way c] Specimen holder
1 The microtome knife is carefully inserted into the knife holder & secured with two
clamping screws.
43
2 Samples are secured in a specimen holder.
There is a knob which is used to clamp the
specimen in place.
3 If the specimen is delicate, it needs to be
encased in a support material.
4 The height of the specimen is regulated by a
lower knob on the specimen holder. There is a lever which locks the specimen
holder in place.
5 The specimen holder needs to be completely withdrawn at the start.
6 The specimen needs to protrude above the top of the specimen holder by 5-10 mm.
7 Its height depends on its size and strength.
8 If it is too high it may bend.
9 If it is too low you may only get a few sections.
10 The knife holder should be fully retracted at the outset.
11 The knife angle is controlled by a
lever on the knife holder.
12 Start with an angle of 10 degrees.
You may need to vary this for
different materials.
13 Section thickness is controlled by a
sliding gauge at the rear of the
microtome.
14 The scale reads in microns. The
thickness is set at 30 microns in the
image. This can be adjusted to
lower settings by unlocking the
gauge and repositioning the scale.
15 In order to cut identical sections, the knife holder must be completely returned to its
starting position at the rear of the slideway. This displaces the gauge completely.
Partial returns produce thinner sections.
16 Sections thicker than 30 microns can be obtained by advancing the knife holder
assembly without cutting a section & retracting it completely. Each time you do
this, the specimens will raised by 30 microns or whatever thickness setting you are
using.
17 This allows you to cut thick and thin sections without changing the thickness
gauge.
18 Position the leading edge of the knife above the specimen holder.
19 Use the knob on the specimen holder to adjust the height of the sample so that it is
very close to the knife edge.
20 Be sure that the two do not make contact at any point!
21 Set the thickness gauge to maximum and pull the knife holder assembly it smoothly
over the specimen until sections are cut. Use the handle on the outside of the knife
holder assembly to move it back & forth.
22 The first sections will be fragmentary and can be whisked away with a wet brush.
23 The wet brush is used to remove good sections by sliding sections onto a slide
containing water, or by getting sections to adhere to it. The latter is not as hard as it
sounds!
24 Sections can be floated in a Petri dish of water or placed directly in water on
microscope slides.
44
25 Best sectioning occurs when the specimen and the knife are wet. This can cause the
knife to rust. Consequently, the knife needs to be dried and oiled ASAP.
26 Once you get reasonably intact sections, place them on a microscope slide and
examine them to see if you are getting the correct Plane of Section.
WARNING - Microtome Knives are EXTREMELY SHARP & Dangerous. They are so
sharp that you can hardly feel them cutting into your hand. Be constantly aware of
the blade'
s location!.
SECTION CUTTING
A solid microtome, such as a base sledge, is best suited to cutting sections of wood
because of its stability and capacity to hold the knife at an oblique angle. A wedge knife
should be used. Sections are easier to obtain from wood impregnated with alcohol/glycerin,
if the surface of the wood is kept wet with glycerin or 70 % ethanol. Sections tend to curl
on cutting and can be flattened with a soft, camel haired brush. Place the section on to a
clean glass slide and mount under a coverslip using Canada balsam as a mounting medium.
Method for soft woods (not embedded)
The sample should be restricted to a maximum face size of 2 cm x 2 cm. Prepare a clean,
smooth surface and paint with a liquid, plastic polymer. This will infiltrate through the
wood which should be dry enough to cut in 20 to 30 minutes at room temperature. With a
sharp, wedge knife sections of 10 to 20 µm should be possible. Thinner sections can be
obtained if the subsequent method for hard wood is used.
Method for hard woods (not embedded)
Sections of hard wood will cut at approximately 10 to 20 µm, depending upon the hardness
of the material and the sharpness of the knife.
Method
1. Maximum block face size 2 cm x 2 cm
2. Boil the wood in distilled water until the wood submerges (this can take up to a few
hours for very hard woods).
3. Dehydrate with 70% ethanol using at least 3 changes over a period of 24 hours.
4. Transfer to a solution of glycerin/70% ethanol and impregnate with at least 3
changes of solution over a period of 24 hours.
WAX METHOD
Suitable for wool, cotton silk, some rayons and most foodstuffs.
Reagent required
Mix together in a heated crucible:
Paraffin wax (melting point 58°C)
= 9 parts
45
Beeswax
Carnauba Wax (vegetable wax)
= 0.5 parts
= 0.5 parts
Method
•
•
•
•
Pour the hot wax mixture over a small piece of the specimen (block face size 1 cm
x 1 cm) positioned in an embedding mould or fibres fixed firmly in an embedding
frame (The embedding frame should be placed on a flat surface such as a glass
plate so that hot wax does not flow from the underside). The slots in the frame, in
which the fibres lie, are sealed with adhesive tape before pouring the embedding
mixture into the mould.
Allow the wax to solidify at room temperature. Avoid rapid cooling as this may
encourage the development of cracks in the hardened block.
Trim the block with a sharp blade then cut sections in the normal way using a
wedge profile knife. Float sections on to warm water to flatten and collect onto a
clean, glass slide.
Remove the wax from the section with xylene then mount the fibres under a
coverslip using Canada balsam as a mounting medium.
PARAFFIN SECTIONING
With a good knife edge and hard, well set and homogenous paraffin wax, sections
of 1 µm are possible if the block face is no larger than 1 cm x 1 cm. In a block of this size
the tissue should occupy approximately 50 % of the surface to be cut. However, the normal
practice since block sizes can be much larger than this is to cut ribbons of sections at 3 to 5
µm. The objective is to produce a ribbon of artifact free, flat sections from which one to
several are selected and mounted on to clean slides.
Method
1. Set the blocks on to a cold surface to harden the face to be cut (a refrigerated cold
plate or ice). Avoid prolonged cooling and very cold surfaces as both can cracks the
surface of the paraffin wax block.
2. Install a sharp, trimming knife in the microtome and set the correct clearance angle,
normally 2 to 5o. (See Knife and cutting angles).
3. Trim a paraffin wax block, with a sharp blade, so that the sides are parallel and 2 to
3 mm of wax surrounds the tissue.
4. Fit the trimmed block into the block holder and orientate so the edge offering least
resistance meets the knife edge first.
5. Advance the block until it just touches the knife edge.
6. Coarse cut the block at 15 µm until the full face has been trimmed.
7. Return the trimmed block to the cold surface for 1 to 2 minutes.
8. Set the advance feed to the desired thickness (3 to 5 µm for most purposes).
9. Remove any debris associated with coarse cutting from the knife edge with alcohol.
(Xylene should not be used as it often leaves an oily remnant on the knife to which
cut sections will stick).
10. Install a fresh, sharp knife in the microtome or move the previous knife to a new,
unused area.
11. Re-install the cold block in the microtome and cut a series (or ribbon) of sections at
the required thickness. Gently breathing upon the sections as each is cut dissipates
46
static electricity, flattens the section and facilitates movement of the ribbon down
the knife. Section compression is minimized by using a sharp knife set to the
correct clearance angle.
12. The ribbon is separated from the knife edge with a moist camel hair brush and
pulled across the surface of a warm water bath (Section expansion will compensate
for the compression caused when cutting).
The temperature of the water should be approximately 10°C below the melting point of
the wax used in the block. Wrinkles in the section can be removed along with small air
bubbles trapped beneath the wax, by careful prodding with a moist camel haired brush or
metal probe (although the latter may damage to the section). Wrinkles usually develop
because different tissue components expand at different rates as the section warms on the
surface of the water.
13. Sections can be separated whilst floating on the water with gentle pressure from the
tips of forceps.
14. Sections are collected on to a clean glass slide. The slide is held vertical and mostly
beneath the surface of the water. The section is apposed to the slide which, when
lifted from the water, draws the section with it.
15. Slides are dried for a minimum of 10 minutes to ensure the section is firmly
attached. A hot plate set just above the melting point of the wax or a hot air oven
are both effective. For delicate tissues more prolonged drying in a hot air oven at a
lower temperature may prove beneficial. Overnight drying at 37°C is necessary for
maximum section adhesion. Slides left in the open for drying will accumulate dust.
All purpose glass slides are 76.2 x 25.4 mm (1" x 3"). Those preferred for light microscopy
are normally 1 to 1.2 mm in thickness and have ground and polished edges to reduce the
risk of injury. Slides with frosted ends are preferred as section/specimen details can be
inscribed with pencil rather than with a diamond stylus, which can create small spicules of
flying glass.
Dry cutting
This technique can be used for a wide range of materials including, wood, paper,
leather, some plastics, varnishes, pigments and metals. Sections obtained always curl or
fray during the dry cutting procedure and it is advisable to use an adhesive polymer or
adhesive foil to keep the specimen together.
METHOD
1. After trimming to a flat surface, a piece of adhesive tape is pressed on to the
surface of the block before each section is cut.
2. The section, attached to the adhesive, can be examined directly or both can be
mounted under a coverslip using a suitable mounting medium.
47
Resin and wax method
This procedure is suitable producing ribbons of serial sections (5 to 6 µm) of all
types of fibres except for those that are very hard or incompatible with the resin (such as
polyamide and polyacryl nitrite). Paper, leather, pigments and foodstuffs can also be
prepared this way.
REAGENT REQUIRED
Mix together in a heated crucible:
Candelilla wax
=5 to 6 parts
Carnauba wax
=1 part
Beeswax
=1 part
Colophony
=1.5 parts
Venetian turpentine
=1.5 parts
METHOD
1. Pour the hot resin/wax mixture over the dry specimen in an embedding mould or fibres
mounted in a metal frame which has been placed in a desiccator. Evacuate the air
carefully to ensure that the embedding medium does not foam. Cool in air and trim to
size with a heated knife.
2. Allow the block to set for approximately 2 hours then cut and float sections on to warm
water to flatten.
3. Collect sections on to clean, glass slides and allow to dry thoroughly.
4. Remove the resin/wax mixture by treating with hot trichlorethylene or carbon
tetrachloride. Both of these substances are extremely hazardous and this procedure
should always be performed under a suitable fume extraction unit. Protective clothing
comprising a long sleeved laboratory coat or gown with elasticised wrist bands, rubber
gloves, safety goggles and a respirator, with canisters suitable for the chemical fumes
should be worn.
5. Mount the section under a coverslip using Canada balsam as a mounting medium.
RESIN METHOD FOR LIGHT MICROSCOPY
This method is applicable to all types of specimens including textile fibres, natural
fibres, woods, paper, leather, plastics, paints and pigments. A rotary microtome fitted with
a glass knife or a base sledge microtome with a wedge shaped knife set obliquely in the
cutting plane are best used for cutting resin sections. The resin is a mixture of methyl and
butyl methacrylate. Stabilisers (usually hydroquinone) normally incorporated into
methacrylate resin to prevent polymerisation during transportation or storage must be
removed before embedding.
48
REMOVAL OF RESIN STABILISER
METHOD
1 Pour the resin to be cleared into a separating funnel of appropriate size.
2 Add (approximately 10% v/v) a 5% to 10% aqueous solution of sodium hydroxide.
3 Shake vigorously for 1 minute. A brown deposit forms and settles to the bottom of
the separating funnel after which it can be drained.
4 Repeat the procedure until the sodium hydroxide remains clear.
5 The methacrylate becomes cloudy after this procedure. It can be clarified by
repeating the procedure using distilled water.
REAGENT REQUIRED
Resin embedding mixture: The exact proportions will vary depending upon the type and
number fibres to be embedded. Between 1% and 2% by volume of accelerator (normally NN dimethyl aniline) is added to the volume of resin to initiate polymerisation.
METHOD
1 Gelatin capsules are suitable as embedding moulds - fibres can be drawn through
the top and bottom of the capsule using a suture needle or material for embedding
can be processed in situ in the capsule.
2 Tie a knot in the fibres at the bottom end of the capsule.
3 Pour the polymer resin into the capsule.
4 Place the capsule lid over the base and pull the fibres tight. Polymerise at 48°C for
5 to 10 hours. Polymerisation at temperatures above 48°C can cause bubbles to
form in the methacrylate.
5 The hardened block is removed by immersing the capsule in water.
6 Trim the block with a razor blade and cut sections using a plane wedge knife or
glass knife.
7
Sections can be removed from the knife edge with a camel hair brush, transferred
to warm water to flatten and then collected on to clean glass slides.
8 Mount sections under a coverslip using castor oil as a mounting medium. The
edges of the coverslip need to be sealed with nail varnish to prevent evaporation of
the mounting medium.
49
Practical 14
STAIN AND STAINING
Stains are a substance which is used to colour the microscopical objects. It is more
rigidly specified and chemically defined brand of the dye. Dyes are comparatively crude
preparations. In other words stains are specified dyes used to make structural details of
cells and tissues more perceivable. A stain is usually certified by the Biological Stain
Commission (Geneva) and bears certification number.
A stain is categorized as acidic or basic stains depending upon whether its
auxochromic groups are acidic (-OH and COOH) or basic (-NH2). Stains are not supplied
as acids or bases. Acid stains are first made into salt using a strong alkali (such as
hydroxide of ammonium, potassium or sodium). Basic stains are salted with acids like HCl
or H2SO4.. Acid stains are, therefore, available as salts of ammonium, potassium or sodium
and basic stains as bromide, chloride or sulphide. In commercially available salt of an acid
stain, most part of the stain is contained in the anion (-) whereas in the salts of a basic
stain, it is in the cation (+). Thus an acid stains is called as an anionic stains and a basic
stains as a cationic stain. When the solution of acid (anionic) and basic (cationic) stains are
mixed, the exchange of ions between two types of heteroionic molecules results into the
formation of neutral stain (e.g. Giemsa stain). Ordinarily, neutral stains are insoluble in
water. Therefore, during staining procedure, they are solubilized in alcohol. Given below
are commonly used stains, their reagents and method of preparation.
1. Safranin
Safranin
Alcohol (95 %)
Water
=1g
= 50 ml
= 50 ml
For aqueous stain it may be prepared as given below
Safranin
Water
=1g
= 100 ml
When ready made stains are available dilute to 1:10 ratio before use
Safranin is suitable for staining lignified or cutinized cell.
2. Aceto-Carmine
1.
It is excellent nuclear stain, suitable for staining pollen grains, chromosomes,
etc. Use 1 % aceto-Carmine; Do not use more than a month old stain
2.
Preparation: Dissolve 45 ml acetic acid in 55 ml water and then add 0.5 g
carmine powder; Heat to boil the mixture for 30 min and then make it cool.
Filter the stain through Whatman No 1 filter paper. At end add one or two drop
50
of Ferric Chloride or Ferric acetate solution. Keep the filtrate in amber colour
bottle.
3.
Caution: Do not stir the stain mixture while filtering
3. Propiono-Carmine
Similar to Aceto-Carmine; Use propionic acid instead of acetic acid while
preparing stain
4. Eosine: Stains the cytoplasm
Eosine
Water
=1g
=100 ml
5. Aniline blue: Also known as China blue, Water blue or Cotton blue. It is good for
cellulose walls. Used for fungi
Aniline blue
Alcohol (95 %)
=1g
= 100 ml
6. Light Green: Also known as Methyl green. It is good for cellulose and lignified cell
walls
Light green
Alcohol (Absolute)
Clove oil
=1g
= 5 ml
= 95 ml
7. Hematoxylins
It is a chromogen derived from logwood Haemotoxylon compechianum. Commonly
employed hematoxylins are i) Heidenhain’s and ii) Delafield’s hematoxylin
i) Heidenhain’s hematoxylin
Half per cent solution of the stain is prepared in warm and distilled water. It is then
stored and closed and dark bottle to ripen at least for four days before use.
ii) Delafield’s hematoxylin
Stock A
Hematoxylin
Alcohol (Absolute)
=1g
= 6 ml
Stock B
Ammonium Alum
Distilled Water
= 10 g
= 100 ml
51
Procedure
Add Stock A to Stock B very slowly.
Expose to light and air for one week and then filter.
Add 25 ml Glycerine and 25 ml Methyl alcohol
Allow to stand uncorked until the colour is darkened. Then filter and keep in
well stoppered bottle.
Allow the solution to ripen before use for a month. Therefore, teachers may
plan well before starting the experiment.
8. Methylene Blue
Solution A
Methylene blue
=1g
Alcohol (Absolute)
= 30 ml
Solution B
Potassium hydroxide 0.001% by wt
= 1000 ml
Mix solution A and B thoroughly
9. Toluidine Blue
Toluidine blue
Acetic acid, 5 ml
Alcohol (Absolute)
Distilled water
= 0.25 g
= 5 ml
= 5 ml
= 100 ml
Dissolve 0.25 g TB, 2 ml absolute alcohol in 100 ml water
10. Fast green
Fast green
Alcohol (90 %)
= 0. 5 g
= 100 ml
11. Erythrosine
Erythrosine
Alcohol (90 %)
=1g
= 100 ml
Or
Erythrosine
Alcohol (Absolute)
Clove oil
=1g
= 5 ml
= 95 ml
12. Crystal violet
Crystal violet
Water
=1g
= 100 ml
52
13. Basic Fuchsin : Good for bacteria and is a strong nuclear stain
Solution A
Basic Fuchsin
Alcohol (95 %)
= 0.3 g
= 10 ml
Solution B
Phenol melted
Distilled water
=5g
= 100 ml
Mix solution A and B thoroughly
STAINING
Staining is a process with which the desired structure is made more discernible
under the microscope. It is physico-chemical phenomenon. A morphologist would stain the
tissue preparation to make its histological details clearer while the main interest of a
biochemist would be to the chemistry and distribution of biosysnthates in a histological
preparation. Staining procedure is flexible and thus a large array of modifications in its
modus operandi is usually available, depending largely on the nature of materials and on
the way of the materials has been prepared i.e. fixed, embedded or sectioned.
TYPES OF STAINING
1. Single staining: Only one stain or dye is used. Common in lower plants staining
where tissues are not differentiated or only a specific structure needs distinction.
2. Double staining: Two dyes are used wherever tissue differentiation is elaborate.
One is acidic and other basic. Unlignified cells are stained by acidic dye and
lignified cells are stained by basic dye. e.g. i) Hematoxylin and safranin, ii)
Safranin and Aniline blue, iii) Safranin and Crystal violet, iv) safranin and fast
green and v) Crystal violet and erythrosine
DOUBLE STAINING PROCEDURE
1. For semi-permanent and temporary preparations
1. The selected sections are transferred from watch glass containing water to another
watch glass containing principal stain (e.g. hematoxylin, safranin, or crystal violet,
etc).
2. The sections are allowed to remain in the stain for some times (4-5 min but you
have to standardize for the tissues or crops).
3. Excess stains are washed repeatedly with water.
4. If de-staining is not complete, section are washed with acid alcohol and then wash
the traces of alcohol with water thoroughly
5. This is followed by counter stain the section (e.g. Safranin or Fast green or
Erythrosine). Transfer the section to a watch glass containing counter stain.
53
6. Since the counter stain act much faster than principal stains, do not keep the section
for prolonged time in the counter stain. Allowed a minute or less than that.
7. Remove the excess stain with 15 to 20 % Glycerine.
8. You will see a section with distinct tissue systems while preserving the colour of
the stain.
9. Mount the section using suitable mountant.
Specific schemes for double staining for temporary mounting
Hematoxylin and Safranin
Safranin and Fast green or Aniline blue
Select a best section & place in 50 % alcohol
Select a best section
Stain with hematoxylin
Stain with Safranin (4-5 min)
Wash with water
Wash repeatedly with water
Wash with ammonia water till stain turns
blue / use tap water if alkaline
Destain with acid alcohol if needed
Wash with water many times
Wash with water
Stain in Safranin dissolved in 50 % alcohol
Destain in 50 % alcohol, wash in ammonia
Mounting in glycerine
Stain with Fast green / Aniline blue (1min)
Wash thoroughly in glycerine
Mounting
2 For -permanent preparations
If an object is to be mounted in Canada balsam and is desired to be stored for a long
time, procedure should include dehydration and clearing.
1.
The section is first stained with principal stain (aqueous e.g. hematoxylin,
safranin, crystal violet, etc).
2.
Wash the section with water till there is no more stain dissolving in water (at
this stage washed water remains colourless)
3.
The dehydration of the section now begins. Pass the section through a graded
series of alcohol. A staining jar is filled with requisite amount of alcohol.
transfer the section into the jar and cover with lid. First grade of alcohol in the
series is 30 %, then 50 %, 70 % and 90 %.
54
4.
When the section passed to 70 or 90 % alcohol, at this stage counter stain is
employed. (e.g. Alcoholic safranin, fast green, erythrosine, etc). These are
mostly prepared in 80 or 90 % alcohol. These stain acts quickly and as such
sections are to be washed immediately after the requisite time is over.
5.
Destaining is done by washing section with 90 % or absolute alcohol.
6.
Finally transfer the section to absolute alcohol. This completes the dehydration
procedure
7.
Clearing now begins with 25 % xylol (25 ml xylol in 75 ml absolute alcohol).
The sections are gradually passed through xylol series of 25%, 50%, 70%, 90%
and finally to pure xylol. Pure xylol is the last stage of clearing.
(Note: If dehydration is not complete pure xylol becomes white or turbid. At this stage,
section should be passed through a reverse series.)
8.
Mount the section in Canada balsam.
NOTE:
Mordants: When some salts are added to the materials before staining, the behaviour of
stain is altered in positive direction i.e. they form an insoluble compound. They are called
mordants
E.g. Sulphate of iron, aluminium, chromium , etc
55
Specific schemes for double staining for permanent mounting
Hematoxylin and Safranin
Safranin and Fast green &
Crystal violet and Erythrosine
Select a section (if necessary use mordant)
Select a section
Stain in hematoxylin (destain mordant)
Stain in aqueous safranin / crystal violet
Wash in ammonia water / tap water
Wash in water until water is colourless
Dehydration with 30 % alcohol
Dehydration with 30 % alcohol
50 % alcohol
50 % alcohol
70 % alcohol
70 % alcohol
Stain with safranin
90 % alcohol
Destain with 70 % alcohol
Stain with fast green / erythrosine
90% alcohol
Destain with 90 % alcohol
Absolute alcohol
Absolute alcohol
Clear with 25 % xylol
Clear or dealcoholizing with 25 % xylol
50 % xylol
50 % xylol
70 % xylol
70 % xylol
90 % xylol
90 % xylol
Pure xylol
Pure xylol
Mount in Canada balsam
Mount in Canada balsam
56
Practical 15
STUDY THE STRUCTURE OF A TYPICAL PLANT CELL-USING ONION
Protocol
1 Take a square epidermal peeling from the onion bulb
2 Mount in water after staining with Safranin /Eosine.
3 Examine the peeling through low and high power of a microscope
4 The following structure can be seen
Observations
•
Cell Wall: Outer layer of a cell, made of cellulose
•
Protoplasm: Consists of nucleus and cytoplasm
•
Cytoplasm: It is hyaline substance with many inclusions as fats,
carbohydrates, oils, water, minerals, acids and plastids, etc. are present
•
Nucleus: Consists of nuclear membrane, nuclear sap, nucleolus and nuclear
network
•
Vacuoles: Are present in the mature cells
•
Tonoplast: Membrane surrounding vacuole is the tonoplast
•
Plasma membrane: Surrounding the cytoplasm is a stiff layer of cytoplasm
which is known as the plasma membrane. it is non-glandular
•
Endoplasm: Between tonoplasm and ectoplasm is present the endoplasm
57
Practical 16
ANATOMY OF MONOCOT AND DICOT PLANT ORGANS
Plant organs differ from one another in their anatomical characters. The characters
are not absolute and exception can always be found. However, uniformity of characters
allows identification of one organ from other. The following are some main points to
remember.
Distinguishing anatomical features in root, stem, leaf and phylloclade
Root
•
•
•
•
•
Exarch xylem
Radial
Root hairs present Collateral
Endodermis always very clear
VB is radial and exarch
Stem
• Endodermis may or may not present
• Vascular bundle conjoint, collateral and endarch
Leaf
•
•
•
Phloem on lower side
Dorsiventrally flattened
Protoxylem towards upper side
Phylloclade
• Dorsoventrally flattened,
• Bundles collateral, endarch,
• Palisade equally developed,
• Stomata on both sides
A key for identification
1. Vascular bundles conjoint, collateral—(bicollateral) and endarch, endodermis not well
developed
----------------------------------------------STEM (2)
1A. Vascular bundles radial and exarch; endodermis conspicuous with casparian strips
----------------------------------------------ROOT (3)
2
a) Epidermis, cortex and vascular tissues well differentiated;
b) Vascular bundles conjoint, collateral-(bicollateral) and open; arranged in a ring
c) Pith well developed.
----------------------------------------------DICOT STEM
58
2A a) Epidermis differentiated, presence of ground tissue;
b) Vascular bundles conjoint, collateral and always closed;
c) Pith not well marked
----------------------------------------MONOCOT STEM
3 a) Protoxylem groups up to six in number
b) Pith small
c) Secondary growth present
----------------------------------------------DICOT ROOT
3A a) Protoxylem groups more than six in number
b) Pith well developed
c) Secondary growth absent
----------------------------------------MONOCOT ROOT
The major difference between monocot and dicot are,
Parts
Root
Stem
Leaf
Monocot
Dicot
More than 6 xylem bundles
(polyarch);
No
cambium;
No
secondary growth ; pith is large and
well developed
• Hypodermis
Present,
sclerenchymatous
Less than 6 xylum bundles (diarch to
hexarch); No cambium but secondary
growth arises; pith is small or absent
• The cells following hypodermis are
not differentiated. They are
parenchymatous and extend from
hypodermis up to the centre of the
axis-known as ground tissue.
• Hypodermis may or may not
present,
if
present,
mostly
collenchyma
• Cortex-Few layers of parenchyma
extend up to VB
• Endodermis: Generally absent
• Pericycle: Present between VB
• Bundles conjoint, closed, arranged
• Bundles conjoint, closed, scattered
in a ring;
throughout the ground tissue.
• Secondary growth is present as
• No secondary growth as there is no
cambium is present
cambium;
• Bundle sheath absent.
• Bundle sheath present
• Pith well marked, but can’t be
• Pith well marked, parenchyma or
distinguished
sclerenchyma
• Number of closed parallel bundles
• Bundle sheath present which may
be sclerenchymatous
• Vascular bundles collateral, open,
variously shaped
• Intraxylary phloem may be
present
• A single mid rib bundle
59
Differences between dicot stem and root
Tissues
Stem
Root
Epidermis
Presence of thick or thin cuticle; Cuticle thin or absent; hairs always
stem
hairs
generally unicellular and simple
multicellular
and
complex;
stomata present
Hypodermis
It may be either collenchymatous Generally absent
or sclerenchymatous
Cortex
Parenchymatous
/ Generally
parenchymatous,
sclerenchymatous
/ chlorenchyma absent, not generally
chlorenchymatous or all the differentiated.
three types of tissues may be
present
Endodermis
Often absent; if present wavy Generally present, a complete ring,
layer mostly represented by cell show thickening-casparian strips
endodermoid cells
Pericycle
A
few
layered
sclerenchymatous
parenchymatous
Vascular
bundles
Many, conjoint, collateral
bicollateral, open and endarch
Pith
Large and well marked
ring, Single layers
/ parenchymatous
ring;
/ A few (2-6), radial, exarch
Small or absent
generally
60
Differences between monocot stem and root
Tissues
Stem
Root
Epidermis
Multicellular / unicellular Hairs always unicellular, cuticle thin /
hairs, cuticle thick, stomata absent, stomata absent
present
Hypodermis
Sclerenchyamtous
Absent
Cortex
Not differentiated, extends
from the hypodermis up to
the centre of the axis,
mostly
composed
of
parenchyma (ground tissue)
Differentiated clearly from the vascular
tissues, extends from hypodermis to
endodermis, composed of parenchyma
mostly
Endodermis
Not present
A well marked out ring like layer
delimiting cortex from the vascular
tissues, casparian strips.
Pericycle
Not marked out
A well definite well marked layer
composed of thin walled cells.
Vascular
bundles
Many in number, conjoint,
collateral, closed, endarch
protoxylem, VB scattered in
the ground tissue
Pith
Cannot be distinguished
Well marked, parenchymatous
61
Practical 17
EXAMINING ANATOMY OF MONOCOT ROOT-MAIZE
WORK TO BE DONE
1. Collect, after washing, fresh roots from pot grown maize seedlings into an open
mouthed bottle containing water or 70 % ethanol
2. Take hand section as per the methods outlined in chapter 8
3. View the section in 4x or 10x and draw the diagram
4. Describe the structure.
5. The table given below may be taken for guidance
Guidelines to describe a monocot root taking Maize as an example
Outline
Epiblema or
piliferous
layer
Cortex
Endodermis
Pericycle
Vascular
Region
Pith
Remarks
Round
This is outermost layer composed of single row of cells, barrel shaped, thin
walled and compactly arranged, provided with number of unicellular root hairs
Occupies large part of the section, several cells /layer deep., consist of thin
walled parenchyma, angular to round in shape with intercellular spaces
In an old root, when epiblema gets disorganized, a few outer layers of cortex
undergo cutinization / suberization and thus outer part of the cortex become thick
walled. The zone of this thick walled cells also known as exodermis. this is a
protective layer which protects delicate internal tissues from outer forces
It delimits the cortex from stele. Cells are barrel shaped, arranged compactly.
Radial and radial tangential walls often shows casparian strips (thickening); A
few cells lying opposite the protoxylem elements thin walled and known as
passage cells
Follows endodermis and consists of thin walled cells, forms a complete ring,
single layer
Xylem and phloem bundles are radial and numerous groups of xylem and
phloem arranged separately on alternating radii, the condition known as
polyarch. In the xylem groups protoxylem situated close to the pericycle, thus the
vascular tissue is developed centripetally and is known as exarch. Xylem
elements consists of tracheids, vessels and parenchyma, while phloem bundle
consists of sieve tube elements, companion cells and parenchyma, in and
between VB thick walled parenchyma is present which is termed as conjuctive
tissue. protoxylem are annularly or spirally thickened, while metaxylem show
considerable reticulate and pitted vessels. Even after considerable maturity
secondary growth does not take place, there being a complete absence of
cambium
In the centre of the axis, well developed parenchyma; in some cases it becomes
thick walled and lignified.
1. Vascular bundles radial, protoxylem is exarch; 2. Undifferentiated and
massive cortex; 3. Unicellular root hairs
--------------------------Therefore the specimen is ROOT
4. Xylem groups show polyarch ; 5) Pith well differentiated ; 6. Complete
absence of secondary growth
------------------Therefore the specimen is MONOCOT ROT
62
Practical 18
EXAMINING ANATOMY OF DICOT ROOT-BENGAL GRAM
WORK TO BE DONE
1. Collect the fresh root from pot grown Cicer arietenum seedlings into an open
mouthed bottle containing water
2. After washing, transfer a piece of root into watch glass containing water
3. Take transverse section (hand section) as per the methods outlined in chapter 8
4. View the section in 4x or 10x and draw the diagram
5. Describe the structure.
6. The table given below may be taken for guidance
Guidelines to describe a dicot root taking Bengal gram as an example
Outline
Epiblema or
piliferous
layer
Cortex
Endodermis
Pericycle
Vascular
Region
Pith
Remarks
Almost circular in transverse section
Composed of single row of cells, thin walled and unicellular root hairs are
present
Undifferentiated, consist of thin walled parenchyma, several cells deep, with
starchy intercellular spaces. Epiblema is short lived in some varieties, after its
death outer layer of cortex become cutinized known as exodermis.
It separates cortex from stele. Cells are barrel shaped, closely packed. Radial and
radial tangential walls shows casparian strips (thickening); A few cells lying
opposite the protoxylem elements thin walled and known as passage cells.
Follows endodermis, thin walled cells, single layer and forms a ring,
Xylem and phloem bundles are radial and exarch; Four groups of xylem and
phloem arranged separately on alternating radii, known as tetrarch. In the xylem
groups protoxylem situated close to the pericycle, thus the vascular tissue is
developed centripetally (exarch). Xylem elements consists of tracheids, vessels
and parenchyma, while phloem bundle consists of sieve tube elements,
companion cells and parenchyma, in and between VB thick walled parenchyma
is present which is termed as conjuctive tissue. protoxylem are annularly or
spirally thickened, while metaxylem show considerable reticulate and pitted
vessels.
Mature cambium appears as a wavy Meristematic layer below the phloem group
and above the protoxylem elements. As a result of secondary growth, primary
xylem elements are pushed towards the centre, where they meet and obliterate
the pith
Very small, occupies the centre of the axis, parenchymatous, hexagonal or
polygonal; intercellular spaces lacking, due to addition of secondary growth.
1. Vascular bundles radial, protoxylem is exarch; 2. Undifferentiated and
massive cortex; 3. Unicellular root hairs
--------------------------Therefore the specimen is ROOT
4. Xylem groups show tetrarch ; 5) Pith very small ; 6. Cambium appear as
secondary meristem
------------------------Therefore the specimen is DICOT ROT
63
Practical 19
EXAMINING ANATOMY OF MONOCOT STEM-MAIZE
WORK TO BE DONE
1. Collect the fresh stem from maize plant into an open mouthed bottle containing
water
2. Transfer a piece of stem into watch glass containing water
3. Take transverse section (hand section) as per the methods outlined in chapter 8
4. View the section in 4x or 10x and draw the diagram
5. Describe the structure.
6. The table given below may be taken for guidance
Guidelines to describe a monocot stem taking Maize as an example
Outline
Cuticle
Epidermis
Hypodermis
Ground tissue
Vascular
Region
Xylem
Phloem
Remarks
Almost round
A thick cuticle is present
Single layered, with stomata occurring here and there, epidermal hairs
absent
Lying below epidermis, two to three layers thick and sclerenchymatous
Extensive, parenchymatous with intercellular spaces. There is no
differentiation into cortex, endodermis and Pericycle
Bundles are numerous, scattered throughout the ground tissue, each VB is
conjoint, collateral, enclosed and endarch. The bundles nearer the
periphery smaller in size than the central ones, each bundle is more or less
surrounded by bundle sheath
Compose of 4 vessels arranged to form a Y shape; Two smaller annular
and spiral vessels are protoxylem while the bigger two pitted vessels and
also some other tracheids and vessels constitute the metaxylem.
Surrounding and just below the protoxylem is a larger water cavity. It is
formed by the breaking down of the protoxylem elements hence known as
lysigenous cavity.
Composed of sieve tubes and companion cells. Phloem parenchyma is
absent
1. Vascular bundles conjoint, collateral and endarch; 2. Undifferentiated cortex;
3. ground tissue present
--------------------------Therefore the specimen is STEM
4. Endodermis and Pericycle are absent; 5) Vascular bundle closed (cambium
absent); 6. VB scattered; 7) Numerous VB; 8) Prominent bundle sheath
------------------Therefore the specimen is MONOCOT STEM
64
Practical 20
EXAMINING ANATOMY OF DICOT STEM-CUCURBITA
WORK TO BE DONE
1. Collect the fresh stem from Cucurbita species into an open mouthed bottle
containing water
2. Transfer a piece of stem into watch glass containing water
3. Take transverse section (hand section) as per the methods outlined in chapter 8
4. View the section in 4x or 10x and draw the diagram
5. Describe the structure.
6. The table given below may be taken for guidance
Guidelines to describe a dicot stem taking Cucurbita sp. as an example
Outline
Cuticle
Epidermis
Cortex
Endodermis
Pericycle
Ground tissue
Vascular
Region
Xylem
Phloem
Pith
Remarks
Ridged and furrowed, five ridges and five furrowed
Present
Single layered, stomata in furrows, multicellular epidermal hairs present
Differentiated in to hypodermis and general cortex. Hypodermis
(collenchyma) 3-4 selves deep, corner angle thick. General cortex follows
collenchyma. It is chlorophyllous (chlorenchyma).
It is inner most layer of cortex. Barrel shaped, contain starch, single
layered.
Crescent shaped sclerenchymatous, 3 or 4 layered; present between
phloem and xylem
Parenchymatous with in which two rings of VB are present
Bundles are bicollateral, arranged in two rings of five each; endarch. The
bundles outer rings are smaller and lying opposite to ridges and the
bundles in inner rings are bigger and correspond to furrows In between
xylem and phloem cambium is present
No definite order of arrangement but protoxylem and metaxylem can be
distinguished
Internal phloem, present on either side of the vascular bundle
Parenchymatous, few intercellular spaces present
1. Well differentiated cortex; 2). VB conjoint, bicollateral, endarch and open
--------------------------Therefore the specimen is STEM
3) VB in a ring; 4) Pericycle distinguishable; 5). Presence of secondary growth
and internal phloem
-----------------Therefore the specimen is DICOT STEM
65
Practical 21
EXAMINING ANATOMY OF MONOCOT LEAF-MAIZE
WORK TO BE DONE
1. Collect the fresh leaves from maize plant into an open mouthed bottle containing
water
2. Transfer a piece of leaf into watch glass containing water
3. Take vertical section (hand section) as per the methods outlined in chapter 8
4. View the section in 4x or 10x and draw the diagram
5. Describe the structure.
6. The table given below may be taken for guidance
Guidelines to describe a monocot leaf taking Maize as an example
Epidermis
Mesophyll
Vascular
Region
Remarks
Both lower and upper surfaces bounded by epidermal layers, uniseriate,
barrel shaped cells, arranged compactly, both layers thickly cuticularised,
stomata present on both epidermal layers, a few large, empty and
colourless bulliform cells present in upper epidermis
Not differentiated into palisade and spongy parenchyma, all the cells
constituting Mesophyll occupy regions extending from upper epidermis to
lower epidermis, cells alike, isodiametric and contains numerous
chloroplast, compactly arranged, leave only a few intercellular spaces
Bundles are numerous, arranged in a parallel series. Size variable,
collateral and closed, distinct parenchymatous bundle sheath surrounds
every vascular bundle, cells of the sheath possess plastids and starch
grains, 9this layer, thus serves as a temporary storage tissue and also
transports the products of photosynthesis to the phloem), a patch of
sclerenchyma each is present above and below the larger vascular bundles
and extends up to upper and lower epidermis respectively, larger bundles
with distinct and more amount of xylem and phloem than the smaller
ones; bundles with xylem on its upper side (towards upper epidermis) and
phloem on its lower side, xylem consists of tracheids, vessels and xylem
parenchyma; phloem comprises sieve tube elements, companion cells and
parenchyma
1. Mesophyll not differentiated into palisade and spongy parenchyma, 2) stomata
present on both epidermis
--------------------------Therefore the specimen is Isobilateral leaf
Note: Most of the monocotyledonous plants show isobilateral leaves. They show stomata on both
of their surface (amphistomatic), though more abundant on lower side. Mesophyll does not show
any differentiation into palisade and spongy parenchyma.
In this case characteristic bulliform cells (also called motor cells) are present in the upper
epidermis. These cells bring about rolling of the leaf by the changes in the turgor pressure and the
rolling is effective in checking the stomatal transpiration. Other xerophytic characters are: thick
cuticle, sclerenchymatous patches and stomata more on lower sides
66
Practical 22
EXAMINING ANATOMY OF DICOT LEAF-MANGO
WORK TO BE DONE
1.
2.
3.
4.
5.
6.
Collect the fresh leaf from mango tree
Transfer a piece of leaf into watch glass containing water
Take vertical section (hand section) as per the methods outlined in chapter 8
View the section in 4x or 10x and draw the diagram
Describe the structure.
The table given below may be taken for guidance
Guidelines to describe a dicot leaf taking Mango as example
Epidermis
Mesophyll
Vascular
Region
Remarks
Both lower and upper surfaces bounded by epidermal layers, single celled
layers, barrel shaped cells, compactly arranged, Upper epidermis covered
with thick cuticle, lacks stomata; lower epidermal covered with thin
cuticle and interrupted by stomata.
Differentiated into palisade and spongy parenchyma
Palisade parenchyma: Just below upper epidermis, regularly arranged
two layers, cells long and tubular, chloroplast present along the radial
walls, compactly and loosely arranged, leave intercellular spaces, layer
interrupted near the large VB by parenchyma, situated just below the
upper and just above the lower epidermis
Spongy parenchyma: The rest of tissue, spongy parenchymatous cells,
small, varied in shapes and sizes, very loosely arranged, enclose smaller
air spaces, a few lead to the stomatal openings, form sub-stomatal cavity,
cells possess numerous chloroplasts along walls
Consist of numerous, small and large bundles, each bundle is conjoint,
collateral and closed, surrounded by parenchymatous bundle sheath, large
VB (e.g. found in the midrib region) possess an extensive bundle sheath,
extends as a parenchymatous mass both toward lower and upper
epidermis, phloem consists of sieve tube elements, companion cells and
parenchyma; xylem made of tracheids, vessels and xylem parenchyma;
metaxylem elements located towards the lower epidermis; while
protoxylem directed towards upper epidermis; phloem situated in a region
of the VB toward the lower epidermis.
1. Mesophyll differentiated into palisade and spongy parenchyma,
--------------------------Therefore the specimen is Dorsiventral leaf
Note: Most of the leaves of dicotyledonous plants are dorsiventral. They grow in
horizontal direction with distinct upper and lower surfaces. This provides more
illumination to upper surface and lesser to the lower surface. Palisade forms a few layers
near the upper epidermis, while spongy parenchyma occurs near the lower epidermis.
67
Practical 23
ECOLOGICAL ANATOMY AND SPECIAL ANATOMICAL FEACTURES
Exercise I
Study the anatomy of the following plants to understand the internal morphology in
relation to ecological adaptation. Check for the presence of key characters given below for
each group viz., hydrophyte, mesophyte and xerophyte
1.
2.
3.
4.
Eichhornia –Hydrophyte-examine root and petiole
Nymphaea-Hydrophyte-Petiole and leaf
Calotropis—Xerophyte-Stem and leaf
Casurina-Xerophyte-Leaf and stem
Hydrophyte
a.
b.
c.
d.
e.
Aerenchyma present
Stomata absent
Cuticle absent
Air cavity present
Reduction of all mechanical tissues
Xerophytes
1
2
3
4
5
6
7
Thick cuticle
Epidermis thick walled
Sunken stomata
Sub-stomatal hairs
Palisade well developed
Mechanical tissue well developed
Transfusion tissues in Casurina
Mesophyte
1.
2.
3.
4.
No sunken stomata
Cuticle moderately thick
Mechanical tissue moderately developed
Intercellular spaces in cortex
Also study the special anatomical features in
•
Phyllode in Acacia moniliformis (Australian Acacia)
•
Phyllocade in Cactus
68
Practical 24
STUDY ON STOMATA IN HYDROPHYTE AND XEROPHYTE
Methods:
There are three methods are available to study the stomata. They are,
1 By using peeling of leaf epidermal layer
2 By using peeling of adhesive gum coated over the leave
3 By taking sectioning
LEAF PEELING METHOD
1 Wash the leaves in water. If needed dip in acetone to remove the chlorophylls
2 Make puncture on the lower or upper epidermis using a forceps and remove a
strip of epidermal layers
3 Stain the epidermal peeling with safranin and
4 Examine under a microscope
5 Count the number of stomata in a microscopic field
ADHESIVE PEELING METHOD
1 Wash the leaves in water. If needed dip in acetone to remove the chlorophylls
2 Spread a thin layer of adhesives /quick fix on the lower or upper leaf surface
3 Wait for a minute to dry the adhesive
4 Then at one end of the leaf surface, lift the adhesive and hold in between thumb
and forefinger
5 Gently pull part the adhesive; it will come as a long ribbon or stripe
6 Examine the ribbon under a microscope for the imprint of stomata
7 Count the number of stomata in a microscopic field
Additional task: Compare the number of stomata on upper and lower leaf surface in
hydrophytes and xerophytes. Deduce the reasons for the difference, if any. For guidance
see an exercise on Ecological Anatomy.
How to work out Stomata Index
Describe terminologies connected with stomata and type of stomata
69
Practical 25
STUDY ON MITOSIS IN ROOT TIPS
Step 1. Collection of root tip
This is the most crucial step in this exercise. The underlying principle is that the
Meristematic cells which are in active cell division have to be collected and preserved after
arresting its life process (i.e. further cell division). Normally root tips detached during
dawn (5 to 7 AM) have more chances of having actively dividing cells. However, this is
tentative and varies with crop. Therefore, by trial and error method you have to standardize
the ideal time for collection of root tip. First survey the available literature and then
according to the local climate find out the suitable period.
Step 2. Pre-treatment
To clear the cytoplasm and hydrolyses the cells pre-treatment is recommended in some
cases. Chemicals like Colchicine 0.5 to 1.0 % in aqueous solution or saturated aqueous
solution Para-Dichlorobenzene are used. The duration of treatment may vary from 3 to 5
hrs at 12-16 0C.
Step 3. Transfer the root tip into a fixative
This is also a grey area. No fixative is standard. However, Randolph’s modified
Navashin fluid and Carnoy’s fluid, FAA are recommended for initial testing.
Step 4: Squash preparation and examination
1 Slides should be perfectly clean. For that immerse the slides in glassware cleaning
solution an hour ahead of your practical. Then clean the slides with running water
and clean and soft cloth or low cost tissue paper
2 Place a small piece (1 mm) of root tips that were collected at appropriate time and
killed (fixed) in appropriate fixatives on the slides
3 Add a drop of water on the specimen and by using the flattened backside of a
needle or pencil crush the root tip by force so that the cell are disintegrated and
nucleus are detached from cells
4 Remove the debris and larger pieces of tissues from the slide with a needle
5 Add a drop of 1% acetocarmine stain over the smeared root cells. (This is a general
recommendation. you may change the stain or modify its concentration according
to your suitability)
6 Place a cover slips over the preparation and if required remove the excess stain by
adding water from one side of the coverslip
7 Gently warm the slide by holding the lower surface of the slide over a sprit lamb
(just few seconds)
70
8 Select the cell which are in metaphase stage (chromosomes are arranged in a line
on the equator and condensed chromosome visible)and observe the chromosome
under high power.
9 Count the number of chromosome and expressed as 2n number.
10 If permanent or semi-permanent slide is required then follow the steps given below.
Step 5. Dehydration before Mounting
After examining the chromosome select one or two best cell. Then mark a circle on
the coverslip just above that particular cell and also on two corner of coverslip extending
the pen marking on the slide also. This will easily guide you to locate the cell after
mounting. Pass the slide through a graded series of Alcohol i.e. 30 %, 50 %, 70 or 90 %.
Remember to always cover the trough containing alcohol series.
Step 6: Mounting
Remove the slide from 90 % alcohol. Air dry the slide for a minute. For semi-permanent
slides use diluted Glycerol. Permanent mounting can be done using DPX
Guidelines to different stages in mitosis
The Cell Cycle
• All cells have come from preexisting cells through cellular division.
• The cell cycle is the period from the beginning of one cell division to the beginning
of the next.
• Somatic cell division involves to main processes: mitosis and cytokinesis; division
of the nucleus and division of the cell, respectively.
• For the most part, in plants cellular division takes place in localized areas called
meristem.
• A cell cycle is divided up into 5 stages
• Interphase is the stage between successive divisions (the end of one to the
beginning of the other)
i. G1: first gap phase – cell growth; synthesis of certain enzymes used in
DNA replication
ii. S: synthesis phase – DNA is replicated in the nucleus; can’t be seen with
a microscope
iii. G2: second gap phase – increased protein synthesis as the cell prepares
to divide.
Mitosis
•
•
•
During mitosis, the nucleus will divide.
All normal cellular activities are suspended.
Each daughter nucleus will have the same number of chromosomes as the parent.
Prophase
• Chromatin (thread-like DNA material) condenses into chromosomes
71
•
•
•
Nuclear envelope and nucleolus disappear
Each chromosome occurs as a double chromo9som (s sister chromatids). These
2 chromatids are sister chromatids because they are exact copies of each other
(recall that DNA replicated during interphase). They are attached to each other
at a central point called a centromere.
Microtubules organize to make the mitotic spindle
Metaphase
• Chromosomes are lined up in the middle plane of the cell
• Mitotic spindle complete
• A spindle fiber from each pole attaches to the centromere of each chromosome.
Anaphase
• Sister chromatids are pulled apart to opposite poles
• Cytokinesis starts
Telophase
• Chromosomes decondense
• New nuclear envelope forms around each set of chromosomes.
• Nucleoli appear
• Spindle fibers disappear
• Cytokinesis ends with the formation of 2 new cells.
o
Cytokinesis is a process of cellular (cytoplasm) division
o
Vesicles begin to congregate at middle point
o
Vesicles fuse together to make 1 large flat vesicle that spans the width of
the cell
o
Their membranes new become the new membrane of each wall
o
Their contents form the new cell wall
Mitosis in onion
72
Practical 26
STUDY ON MEIOSIS IN FLORAL BUDS
Step 1. Collection of floral buds
Similar to earlier exercise on mitosis, this is the most important step in this
exercise. The underlying principle is that the gametophytes-PMC or EMC (i.e. pollen
mother cells or embryo mother cells) which are in active cell division (reduction division)
have to be collected and preserved after arresting life process in them (i.e. further cell
division). Normally floral buds of Poaceae plants which produce numerous floral buds in
an inflorescence are ideal for this exercise. Immature buds are detached from plants at
opportune time and can be found out by trial and error method.
Step 2. Transfer the root tip into a fixative
This is also a grey area. No fixative is standard. However, Randolph’s modified
Navashin fluid and Carnoy’s fluid, FAA are recommended for initial testing.
Step 3: Squash preparation and examination
1
Slides should be perfectly clean. For that immerse the slides in glassware
cleaning solution an hour ahead of your practical. Then clean the slides with
running water and clean and soft cloth or low cost tissue paper
2
It is easy to study the process of meiosis in PMC than EMC as the later
produces only one PMC per flower while it is up to four in each anther in the
former. Each PMC also produces many pollen grains. Hence anther is taken for
this exercise
3
Remove the anther from floral bud. Add a drop of water on it and by using the
flattened backside of a needle crush the root tip by force so that the cell are
disintegrated and PMCs are detached from cells
4
Remove the debris and larger pieces of tissues from the slide with a needle
5
Add a drop of 1% acetocarmine stain over the smeared root cells. (This is a
general recommendation. you may change the stain or modify its concentration
according to your suitability)
6
Place a cover slips over the preparation and if required remove the excess stain
by adding water from one side of the coverslip
7
Gently warm the slide by holding the lower surface of the slide over a sprit
lamb (just few seconds). This step is optional
8
Select the cell which are in Diakinesis stage of meiosis I (chromosomes
attained maximum condensation hence visible) and observe the chromosome
under high power.
9
Count the number of chromosome and expressed as 2n number.
73
10
If permanent or semi-permanent slide is required then follow the steps given
below.
Step 4. Dehydration before Mounting
After examining the chromosome select one or two best cell. Then mark a circle on
the coverslip just above that particular cell and also on two corner of coverslip extending
the pen marking on the slide also. This will easily guide you to locate the cell after
mounting. Pass the slide through a graded series of Alcohol i.e. 30 %, 50 %, 70 or 90 %.
Remember to always cover the trough containing alcohol series.
Step 5: Mounting
Remove the slide from 90 % alcohol. Air dry the slide for a minute. For semipermanent slides use diluted Glycerol. Permanent mounting can be done using DPX
Theoretical background of Meiosis
In the process of sexual reproduction, sex cells called gametes unite to form a
zygote. The zygote will then undergo mitosis to form the new individual. However, in
order for the original number of chromosomes to be maintained across generations,
gametes need to have half the normal complement of chromosomes. Meiosis is a special
type of mitosis that reduces the number of chromosomes in the daughter cells. Meiosis
produces haploid cells form diploid cells. Meiosis separates homologous chromosomes
into different cells and results in 4 haploid cells from 1 diploid cell.
Meiosis I
•
•
Starts with a regular diploid cell
Chromosomes duplicated during interphase
Prophase I:
o Homologous chromosomes arrange along side each other (called synapsis);
1 set of sister chromatids come from father and other set from mother).
o Crossing over – homologous chromosomes exchange segments. (type of
genetic recombination)
o Spindle forms; nuclear membrane and nucleoli disappear.
Metaphase I – sets of homologous chromosomes line up at middle plate
Anaphase I – The homologous chromosomes are pulled apart to separate poles (not the
sister chromatids of each homologous chromosome)
Telophase I – New nuclear envelopes appear, etc., and cytokinesis. Note that the resulting
cells are now haploid, although the chromosomes are still duplicated.
74
Meiosis II
Interphase II – this stage is very brief and doesn’t even exist in some organisms. DNA is
not replicated this time.
Prophase II – also brief because the chromosomes never completely uncoiled. Nucleus
disappears, chromosomes condense and spindle forms.
Metaphase II – Chromosomes line up at middle plate.
Anaphase II – sister chromatids are pulled to opposite poles
Telophase II – nucleus reorganizes itself; cytokinesis occurs. Note each resulting daughter
cell does not have any duplicated chromosomes.
Differences between Mitosis and Meiosis
Mitosis
Occurs in body cells
Mitosis I is a division event
Two 2N daughter cells formed
Daughter cells same as parent
Meiosis
Occurs in reproductive organs
Meiosis II is a division events
Four N daughter cells formed
Daughter cells genetically different
75
Practical 27
STUDY ON POLLEN FERTILITY
1 Collect the anther or pollen grain just before anthesis / dehiscence
2 Slides should be perfectly clean. For that immerse the slides in glassware cleaning
solution an hour ahead of your practical. Then clean the slides with running water
and clean and soft cloth or low cost tissue paper
3 Fresh anthers from buds are placed in the centre of slide.
4 Crush the anthers with scalpel or just using another slide to liberate pollen grains
5 Remove the debris and larger pieces of tissues from the slide with a needle
6 Add a drop of 1% acetocarmine stain over the smeared root cells. (This is a general
recommendation. you may change the stain or modify its concentration according
to your suitability; use propionocarmine for bhendi, Iodine potassium iodide for
rice, etc)
7 Place a cover slips over the preparation and if required remove the excess stain by
adding water from one side of the coverslip
8 Count the number of fully stained pollen grains and half stained or non stained
pollen grain in a microscopic field.
9 Change the view of microscopic field and again counts the same.
10 Repeat the above step for 5 to 10 times and find out the average in each category.
Count
No
(i)
1
2
3
…
Mean
Number
of
stained pollen
grains in a
microscopic
field
(ii)
Number of unstained /
half stained pollen
grains
in
a
microscopic field
Total
(ii+iii)
(iii)
(iv)
11 Find out the fertility percentage by dividing column 1 by column iv and then
multiply by 100
12 The underlying principle is fertile pollen takes stain while sterile or abnormal
pollen do not.
76
Practical 26
CYTOCHEMISTRY & STAIN SPECIFICITY
Histochemistry or cytochemistry deals with the identification of metabolites in
tissues using stains or similar labels that bind the substrate specifically upon contact. In a
typical histochemical reaction, the dye-substrate binding is in stoichiometric proportions,
thus even a quantification of the substrate can be carried out by evaluating the intensity of
Localisation reaction. The histochemical technique has a definite edge over the
biochemical characterisation as the former does not dislocate the metabolites during the
identification reactions, so that the distributional pattern of metabolites in the tissues can in
addition be visualized. Most of the stain are specific in reaction and are purposefully
employed for definite structure or substances. The list below gives some indication about
usefulness of a stain. Histochemistry involves both physical and chemical process. Specific
stains are employed to distinguish different parts as well the location of different chemicals
in the tissue. The following table will give rough idea about this physio-chemical process.
.
#
Specificity /Tissue / Purpose
Suitable Stains
1
Anatomical figure
Aniline blue, Erythrosine
2
Callose
Aniline blue
3
Cellulose cell walls
Aniline blue, Delafiield’s hematoxylin, fast
green, light green (Methyl green), Congo red
4
Chitinous substances
Safranin
5
Chromosome
Acetocarmine,
Hematoxylin,
Iron
hematoxylin, Methyl green, Safranin, Basic
Fuchsin
6
Cuticularised cell wall
Crystal violet, Erythrosine, Methylene blue,
Safranin
7
Cytoplasm
Aniline blue, Eosine (yellow), Fast green,
Light green, Methyl orange, Hematoxylin
8
Epidermal structures
Basic Fuchsin
9
Lignin
Crystal violet, Safranin, Phloroglucinol plus
dil. HCl, Methyl green (Light green)
10
Mitochondria
Crystal violet
11
Mucilagenous Structure
Crystal violet, Iron hematoxylin, Methylene
blue.
12
Plastids
Crystal violet and Iron hematoxylin
13
Proteins
Safranin
77
14
Suberin
Safranin
Immense scope and versatility of the histochemical techniques have been
undervalued probably due to the near non-availability of text books describing both the
practical modalities and chemical bases of localisation reactions. In this following section
we have presented some useful histochemical exercises with alternative protocol for each
histochemical
Note:
1. For histochemical studies use only distilled water for washing, stain preparation,
etc.
2. It is also recommended to use new slides or well cleaned slides
3. Sometimes the stains stain substrates unspecifically. Therefore it is strongly
recommended to go for a suitable ‘control exercise’ simultaneously to compare
localization of chemical under investigation. Therefore, at the end of each exercise
the procedure for making control sample is given.
78
Practical 27
LOCALISATION OF CELLULOSE / MUCILAGE/CUTIN/SUBERIN / LIGNIN
Method: I2, KI, ZnCl2 staining
Stain preparation: ZnCl2 -30 g; KI-0.5g; I2-0.89 g; H2O-14 ml. First dissolve KI in
distilled water and then I2 and ZnCl2 successively. Keep the stain in dark.
or try this combination : Dissolve ZnCl2 -150.6 g and KI-48.2 g in 50 ml water; Then
saturate the solution with I2-3 g.
Principle
IKI will stain starch blue-black to orange depending on the type of starch present. It
will also stain nuclei a golden color. Cell walls also stain light yellow with IKI. When a
drop of Sulphuric acid added to the stain / tissue it hydrolyses the cellulose into dextrin and
breaks interpolymeric hydrogen links. Iodine, then, penetrates into loosened cellulosic
micelles and dextrin, and produce dark blue.
Protocol
Place several sections on a slide
Flood with IKI
It is frequently not necessary to remove IKI before viewing the sections. Apply a
coverslip and wait a few minutes.
Observation and Interpretation
Blue purple
Brown
Violet purple
Violet
Yellow
Yellowish
Yellow brown
: indicate STARCH
: LIGNIN
: CELLULOSE
: MUCILAGE
: CUTIN
: SUBERIN
: PROTOPLASMIC CONTENT
Specific test for Starch
1. Starch IKI Method: To prepare stain first dissolve 2 g of KI in 100 ml water and then
dissolve 0.2 g (or 1 g) iodine in the KI solution. Hydrated the section of the materials
fixed in FAA or neutral formaldehyde and then place in IKI solution for 10 min.
Wash in water and mount in glycerol: Iodine KI mountant (1:9). Old starch appears
blue to black, whereas newly formed starch appears red to purple
79
CONTROL
Alpha amylase digestion for starch: This test is to digest branched chain polymers or
amylopectin. Incubate the hydrated section in 0.5 w/v solutions of alpha amylase in
0.004 M acetate buffer, pH 5.5 for 3 hrs at 37 0C. To prepare alpha amylase dissolve
0.29 ml of glacial acetic acid in distilled water to make 100 ml solution A. Dissolve
0.288 g of anhydrous sodium acetate in water to make 10 ml solution B. Mix 1 ml
Sol. A with 1 ml Sol. B and make the volume to 100 ml. Adjust pH to 5.5.Dissolve
0.5 g alpha amylase to this buffer and make the volume to 100 ml
Specific test for Chitin
Calcium chloride Test: To prepare stain first dissolve1.25 g of KI in 10 ml water and then
dissolve 125 m g iodine flakes and 40 g of CaCl2. Stain for 15 min in the staining
solution and then mount in the same solution
CONTROL
Digestion with acetic acid: Put the slides in 2 % solution of acetic acid for 5-10 min. after
autoclaving with 23 M KOH at 15 psi, 121 0C. Then go for calcium chloride test.
80
Practical 28
LOCALISATION OF INSOLUBLE POLYSACCHARIDES
Method: Periodic Acid Schiff’s (PAS) reagent test
Stain preparation:
Schiff’s reagent is prepared by dissolving 1 g Basic Fuchsin in 100 ml of boiled water.
After cooling add 0.5 g sodium or potassium meta bisulfite and 10 ml 1N HCl or 100
ml of 0.15 N HCl. Shake the mixture for 2-3hrs and leave it overnight in dark. By the
next day the solution must turn straw yellow due to impurities. To remove this, add
30 mg fresh activated charcoal and shake for 5 min. Filter and keep the colourless
solution in a coloured bottle in the refrigerator. The optimum pH must be 2.3.
Buffered SO2-Water: Dissolve 0.4 g Glycine, 0.3 g NaCl2 and 0.31 ml conc HCl in
distilled water’ make up the volume to 90 ml. Adjust pH to 2.28. Add 10 ml of 15
% w/v sodium bisulphate and mix thoroughly.
0.8 % Periodic acid: Dissolve successively 0.164 g of anhydrous sodium acetate and 0.8 g
of crystalline periodic acid in enough water and make the volume to 100 ml
Protocol
Place the deparafinised section in 0.8 % periodic acid at 40 0C for 10-15 min
Wash in running water for 10 min
Place in Schiff’s reagent for 15-20 min
Remove excess stain by flooding with water
Give three changes in Glycine HCl buffered sodium bisulphate for 1 min each or
transfer the section into 20 % Sodium metabisulphite for 1-2 min
Wash again in running tap water for 5-10 min
Dehydrate in 95 % ethanol, ethanol 1, 2 and mount in DPX
Indication: Polysaccharides including chitin and callose stain intense purplish red colour.
Sometimes unsaturated lipid also stains positively.
CONTROL: Omission of the oxidation in periodic acid may be used as control for
polysaccharides.
81
Practical 29
LOCALISATION OF CELLULOSE
Method: Methylene Blue staining
Stain preparation: Dissolve 1 g Methylene blue dye in 100 ml distilled water and
filter through Whatman No 1 filter paper
(or)
Stock A: 0.3 Methylene Blue g in 30 ml acetic acid
Stock B: 0.01% KOH by weight in 1000 ml
Mix A & B
Protocol
Stain fresh hand cut section for 5 min or longer
Remove excess stain by flooding with water
Mount in Glycerol
Observation and Interpretation
Deep blue colour
Greenish blue
indicate
Pure cellulose
Cellulose mixed in other chemical
CONTROLS
1 Enzymatic extraction: Cellulose may be dissolved by enzymatic extraction. Incubate
the hydrated and cellodin coated sections in cellulose solution at 20-250C for
overnight and then follow the usual staining procedure as outlined above. To prepare
the enzyme solution first prepare 0.005 M phosphate buffer at pH 5.8 by dissolving
0.007 g disodium hydrogen phosphate dihydrate and 0.0627 g potassium dihydrogen
phosphate to a final volume of 100 ml and adjust pH to 5.8. Add 4 g crystalline
cellulose in enough of this buffer for making 100 ml solution
82
Try the other methods
1. Iodine Sulphuric acid method: Prepare Iodine-potassium iodide (IKI) solution by
dissolving 2 g potassium iodide and 1 g of iodine flakes in distilled water to make
100 ml solution. Place the hydrated section of the materials fixed in FAA or 4 %
neutral alcohol in the IKI stain for 15 min. Place a coverslip and then add a drop of
H2SO4 at the side of coverslip and let it diffuse under the glass. Cell wall containing
cellulose stains dark blue; lignin appears orange yellow. Sometime Hemicellulose
gives positive reaction.
83
Practical 30
LOCALISATION OF REDUCING SUBSTANCES
Method: Fehling’s Reagent Test
Protocol:
Flood the sections with a 1:1 mixture of Fehling’s solution A and B
Gently heat the slide
Observation and Interpretation
BROWN DEPOSIT OF METALLIC COPPER
indicate
Reducing substances
Try the other methods:
1. Silver Nitrate reduction Test: Treat section with solution comprising equal
volumes of 0.2 N silver nitrate, 2N ammonium hydroxide and 1 % sodium hydroxide
or 10 5 aqueous silver nitrate. In both cases, any reducing substances present in the
tissues should lead to the formation of a black metallic silver precipitate. The test is
employed for ascorbic acid.
2. Ferric-Ferricyanide Reaction: Treat the section with mixture of 1 % ferric chloride
and 1 % potassium ferricyanide in 2N acetic acid (pH2.25) for 5-10 min. In the
presence of reducing substances, excess ferric ions will be reduced to ferrous state,
leading to formation of ferro-ferricyanide which is blue in clour.
Caution: Be wary of potassium ferricyanide. It is poisonous. Heed all precautions for
the proper use of the chemicals.
84
Practical 31
LOCALISATION OF PECTIC ACID/LIGNIN/TANNIN
Method: Toluidine Blue staining
Principle:
Toluidine Blue is a metachromatic (many colors) stain, and stains lignified walls
blue-green. Unlignified walls with more pectin stain cherry red. However, if you overstrain (too long) with Toluidine blue, everything will be blue.
Stain Preparation: Dissolve 0.25 g TB, 2 ml acetic acid, 5 ml absolute alcohol in 100 ml
water
Protocol
Section cleared in dilute chloral hydrate (2 parts water: 1 part chloral hydrate).
Dried using blotter paper before staining
Wash the section in distilled water
Add several sections to a drop of water on a slide. Add 2-3 drops 0.05 % aq.
Sol. of Toluidine blue for 10-60 seconds
Mount in Water
Observation and Interpretation
Pinkish / purple
Green / Greenish blue / Bright blue
indicates
Pectic acid
Lignin & Tannin
Caution: Toluidine Blue is hard to get out of clothing, so use it carefully and clean up any
spills with lots of water. In addition, it is poisonous, so avoid getting it on your skin as
much as possible. Wear surgical gloves to protect your hands. Be sure to wash your hands
well if they become stained.
85
Specific test for Pectin
1. Ruthenium red Method: Hydrate the section (if it is microtome cut section) and coat
with celloidin. Place the section in 3:1 absolute ethanolic HCl mixture for 24 hrs and
then in Ammoniua solution for 2-5 hrs. Immerse the section in 0.02 % aqueous
Ruthenium Red, keep in dark for 10-15 min. Pectic materials stain red
CONTROL
Enzymatic extraction: Pectin may be dissolved by enzymatic extraction. Incubate the
hydrated and cellodin coated sections in 1 % pectinase solution at 370C for overnight
and then follow the usual staining procedure as outlined above. To prepare the
enzyme solution first prepare 0.005 M phosphate buffer at pH 5.8 by dissolving
0.007 g disodium hydrogen phosphate dihydrate and 0.0627 g potassium dihydrogen
phosphate to a final volume of 100 ml and adjust pH to 5.8. To this add 1 g pectinase
in enough of this buffer for making 100 ml solution
Specific test for Tannin
Ferric Chloride Test: Tannins are complex polyphenolic hence readily react with Ferric
ions to yield blue to green colour complex. Flood the section with 10 % aqueous
ferric chloride or 10 % Fecl3 (or Ferric sulphate) in 95 % ethanol for 10-20 min.
Materials containing phenols and Tannin will stain bluish.
86
Practical 32
LOCALISATION OF LIGNIN
Method: Phloroglucinol-HCl staining
Principle:
Lignin is a phenyl propane polymer. Phloroglucinol-HCl stains are colorless until
it reacts with lignin or suberin, the reaction is known as Weisner’s reaction; at the end
lignified and suberized cell walls will stain red-orange.
Prior requirement:
Section should be cleared in dilute chloral hydrate (2 parts water: 1 part chloral
hydrate) and then dried using blotter paper before staining
Protocol
Stain section in 1 % Phloroglucin prepared either in water or 95 % ethanol for 5 min
Add coverslip. Then add two to three drops 50 % HCl at the side of cover glass or immerse
the section for 1 min in HCL
Mount in Glycerol
Caution:
Lignin appear red colour
1. Phloroglucinol contains 20% HCl. Consequently, clean up any spills, especially on
the microscope stage, and avoid getting this on yourself.
2. Weisner’s reaction (phloroglucinol-HCl) may take several minutes and works best
without a coverslip. If you are going to use several stains or look at several
specimens, stain with Phloroglucinol first, and set these aside until you have
finished other things.
CONTROL
Ethanolysis: Prepare 3 % HCl (8.5 ml in 91.5) in 95 % ethanol. Boiling of the section in
this solution solubilizes lignin by a combination of solvolysis and depolymerization.
Then go with above staining procedure
87
Practical 33
LOCALISATION OF PHENOLS
Method: Toluidine Blue Test
Reagents: Toluidine Blue ‘O’ & 0.1 M Acetate buffer at pH 4.4
(To prepare Acetate Buffer make Stock Sol. A-Mix 11.55 ml of Acetic acid in 1000
ml water; B-Dissolve16.4 g of sodium acetate in 1000 ml; Mix 30.5 ml stock sol. A with
19.5 ml Stock B. make up the volume to ml. this will give acetate buffer of pH 4.4)
Principle:
Toluidine Blue is a metachromatic stain. In strong acidic medium it reacts with
phenolic compounds and stain greenish blue
Protocol:
Stain the sections with Toluidine Blue O in 0.1 M acetate buffer
Greenish blue to Green colour
indicate
Phenolic substances
Caution: Check the pH of acetate buffer since colour persists only at low pH
Try the other methods:
1. Vanillin Test: Sections are dry heated on slides at 150 oC for 5 min before treating with
a drop of fresh saturated solution of (alcoholic) vanillin. Add several drops of conc
HCl. Wash the section and mount. A red colour is produced when aldehyde group in
the vanillin condense with phenols in the tissue.
2. Millon’s Reagent Test: Heat section in an acidified 5 % aqueous solution of mercuric
sulphate for 10 min at 40 oC followed by the addition of 0.5 % Sodium Nitrate.
Coloured Nitrosos derivatives of any phenol contained in the tissue then become
evident.
88
Practical 34
LOCALISATION OF CUTIN AND SUBERIN
Method: Sudan III or IV staining
Principle:
Sudan IV stains suberized cell walls and oil in cells. The stain is dissolved in alcohol.
When the specimen is rinsed with water, waxy and oily materials which have taken up the
stain will remain red or orange, other areas are colorless.
Stain preparation: Add 10 mg of Sudan III dye in 5 ml of 95 % ethanol or rectified spirit.
add 2 g of glycerol. Keep the reagent bottle tightly closed
Protocol
Place several drops of stain on the slide and add sections to it.
The alcohol evaporates rapidly, so it is best to add a coverslip right away.
It takes a few minutes to hrs for the stain to work
(so you will need to add stain periodically to the edge of the coverslip to prevent the
formation of air bubbles)
Wash the stain with 50 % ethanol.
Look for red-orange areas
Red colour
CUTIN / FAT /OIL GLOBULES
Reddish Brown
SUBERIN
Note: Cytologically, cutin and suberin behaves similarly. Cutins are extra cellular
substances, forming major component of plant cuticle. Being esterified with fatty acid it
behaves like lipid. Suberins are secondary cell wall substances occurring inside the tissues
unlike cuticle which is found outside. This characteristic of suberin often used to
distinguish them beforehand from cutin.
89
Practical 35
LOCALISATION OF PROTEIN
Method I: Mercury Bromophenol Blue Method
Principle: Proteins are amphoteric compounds. A protein becomes positively charged at a
pH below the isoelectric point whereas negatively charged (anionic) at a pH which is
higher than isoelectric point. In MBB staining process, first tissues protein is brought
in contact with the acid stain. Rinse in 0.5 %acetic acid brings pH around tissue
protein down so that all proteins, whose isoelectric point lies well above the pH of
0.5 % acetic acid, become positively charged and form electro covalent salts with
anionic bromophenol blue.
Stain preparation: Dissolve 10 g Mercuric chloride and 100 mg of bromophenol blue in
100 ml distilled water or in 100 ml of rectified spirit or 2 % aqueous glacial acetic
acid.
Protocol
Hydrate the section of the materials fixed in neutral buffered Formaldehyde or FAA
Stain the section in the staining mixture for 15 min at room temperature.
Rinse for 20 min in two changes of 0.5 % aqueous acetic acid and in water
Give a tip in absolute tertialry butanol
Clear in xylene
Mount in DPX.
Proteins appear blue.
CONTROLS:
1. Deamination: Prepare the deamination mixture by dissolving 15 g Sodium nitrite in
water and then mixing 0.75 ml of acetic acid into it to make a total volume of 100
ml. Place the section ion the mixture at room temperature for 1-24 hrs.
2. Sulphation: Treat the ethanaol dehydrated section in sulphation mixture consisting of
1:3 mixtures of conc. H2SO4 and acetic acid in volumetric proportion, 3-5 min at
room temperature. This treatment blocks free amino and hydroxyl groups.
90
Try the other staining methods:
1.
Acid Fuchsin Method: Stain the sections with 0.005 % acid fuchsin in 1 %
aqueous acetic acid for 10 min. Protein stain red. This is a useful method to
discriminate between phenols and aromatic amino acids, since the later only stains.
2.
Potassium Ferrocyanide + FCl3 staining: Immerse the fresh section in Stock A
for 1 hr. Rinse immediately with 60 % aqueous acetic alcohol. Add a few drop of
Stock B solution. Protein appears blue. (Stock A-Dissolve 0.8 g Potassium
ferrocyanide in 100 ml of glacial acetic acid. Stock B-Prepare 5 % aqueous Ferric
chloride solution)
3.
Coomassie Brilliant Blue staining: Stain the sections with 0.25% w/v CBB stain
in 7 % aqueous acetic acid for 3 min. Dip the slide for a while in 7 % aqueous
acetic acid to remove excess stain. Blot and air-dry the slides. Mount in 5 % acetic
acid in glycerol. Protein appear in violet colour
4.
Chloramine-T Schiff’s Reaction: Hydrate the tissues fixed in acetic ethanol and
wash it in distilled water. Immerse sections in 1 % solution of chloramines T in 0.1
M Phosphate buffer (pH 7.5) for 6 hrs at 40 oC. Then rinse in distilled water and
transfer to dilute solution of Sodium thiosulphate for 1-2 minutes. Stain in Schiff’s
reagent for 30 min. Wash in distilled water and dehydrate by giving two changes
in t-butanol; pass through xylele before mounting. (Schiff’s reagent is prepared by
dissolving 0.5 g Basic Fuchsin and 0.5 g sodium metabisulfite in 100 ml 0.15 N
HCl. Shake the mixture for 2-3 hrs. Add 30 mg fresh decolourizing Charcoal and
shake for 5 min. Filter and keep the colourless solution in a coloured bottle in the
refrigerator. To prepare Phosphate Buffer (pH 7.5) add 16 ml of Stock A with 84 l
of Stock B and dilute to 200 ml. Stock A: 0.2 M Sol. of Monobasic Sodium
Phosphate (27.8 g in 1000 ml); Stock B 0.2 M Solution of Dibasic Sodium
Phosphate (53.65 g of NA2HPO4. 7H2O or 71.7 g of Na2HPO4.12H2O in 100 ml)
5.
Alkaline Fast Green Method: This test is meant for basic protein i.e. Histones.
Hydrate the tissue prepared in neutral buffered Formaldehyde. Incubate the
hydrated section with 5 % W/v aqueous trichloroacetic acid, in 0.01 M phosphate
buffer at pH 8.0 at 60oC for 80 to 90 minutes. Give 4-5 changes in distilled water,
and then in 70 % ethanol twice allowing 10 min in each change. Again rinse in
water. Stain for 30 minutes in Fast Green solution (50 mg Fast Green FCF in 45
ml 0.01 M phosphate buffer). Give three changes in distilled water and dehydrate
by immersing in rectified spirit. Pass through 50 % ethanolic xylene, xylene 1,
xylene2, and mount. Basic proteins stain green to bluish green. (To prepare 0.01
M phosphate buffer at pH 8.0 dissolve 0.172 g disodium hydrogen phosphate
dihydrate and 0.0042 g potassium dihydrogen phosphate to a final volume of 100
ml and adjust pH to 8.0. (Instead of phosphate buffer try HCl-Borate buffer (pH
8.0). For this 0.05 g of the dye is dissolved in 450 ml of distilled water. To this 50
ml of HCl-Borate buffer (pH 8.0) is added. The stain is stored in 500 ml stopper
bottle after adding a crystal thymol, final pH being 8.1 to 8.2)
91
LOCALISATION OF LIPIDS
Method: Sudan Black B Methods
Principle: Sudan Black B like other Sudan dyes behaves as a solute and enters into
solution-phase with lipoidal substrate, thus staining them. It is neutral dye.
Stain preparation: To prepare acetylated Sudan Black B, dissolve 1 g stain in 100 ml
diethyl ether. Filter and heat the filtrate to boiling under a reflux condenser and then
add 0.5 ml acetic anhydride and 20 ml solvent ether (diethyl ether). Boil under the
reflux condenser for 20 min. cool and filter. Extract the filtrate repeatedly with cold
water until the water layer is neither coloured nor appreciably acidic when tested
with ph paper. Put the ethereal layer into a China dish and evaporate ether. The Black
solid residue is acetylated Sudan Black B. It should have a metallic luster.
Protocol:
Use fresh section either hand cut or paraffin cut
Rinse briefly in 70 % Ethanol
Stain for 5 min in either in saturated solution of Sudan Black B in 70 % aqueous
ethanol (or propylene glycol) or acetylated Sudan Black B (more preferred)
Rinse in 70 % ethanol to remove excess stain
Wash in water and Mount in glycerol-gelatin
Lipid stains bluish black
CONTROL
Immerse in acetone at 600C for 1 hr. This extracts all lipids except the
phospholipids. Reflux for 30 min in three changes in 2:1 v/v mixture of chloroform with
methanol at room temperature. Treat with 3:1 mixture of solvent ether and ethanol and
ethanol for 3 hrs in 3 changes of 1 hr each at 60 0C. Immerse in pyridine at 60 0C for 1-2
hrs, or even overnight, and then wash in running water for 2 hrs. Immerse in benzene at
room temperature for 1 hr. treat with detergent like Triton-100, Tween 80 for 15 min at
room temperature and then wash in water. This treatment removes any bound lipids. Then
follow the usual staining procedure.
92
Try the other staining methods:
Nile Blue Method: Treat freshly sectioned tissues in 1 % Nile Blue at 37 oC for 1 to 2 min.
Then differentiate in 1 % acetic acid at 37 oC for 1 to 2 min. Wash in distilled water
and mount in glycerine-gelatin. Natural lipids will stain red while free fatty acids and
phospholipids will stain blue.
93
Practical 37
LOCALISATION OF NUCLEIC ACIDS (DNA and RNA)
Method: Feulgen method for DNA
Principle: Treatment of Basic Fuchsin with a reducing agent (SO2 or H2SO3) produces a
colourless dye which is readily oxidizable, especially by aldehyde and ketone
groups. Splitting of purines from DNA by dil. HCl releases aldehyde groups or
ketone de novo in the furanose sugar moiety of DNA and react with stain.
Stain preparation
See exercise on Polysaccharides
Protocol
If the section is cut through microtome then coat the section with celloidin (dip in 0.1
% celloidin in 1:1 ethanol ether, after dewaxing in xylene) and hydrate them
Hydrolyse section in 5 N HCl (43.65 ml conc. HCl in water and make the volume to
100 ml) at 25 oC for 1 hr or Feulgen hydrolysis with 1N HCl at 60 oC for 20-30 min
Wash in running water for 10 min
Place in Schiff’s reagent for 15-20 min
Remove excess stain by flooding with water
Transfer the section into2-5 min in Buffered SO2 –water or in 20 % Sodium
metabisulphite for 1-2 min
Wash again in running tap water for 5-10 min
Dehydrate (give three changes in absolute tertiary butanol and two changes in xylene)
Mount in DPX
Indication: Structures containing DNA are stained violet or purple
94
Dos and don’ts:
1.
2.
3.
4.
Do not prolong the treatment of Schiff’s agent with charcoal
pH of Schiff’s reagent must be 2.3
Excess of deficiency of SO2 decrease the colour reaction
Treatment of stained section with water does not remove stain but do it
with tBA
5. Store the slide in dark, otherwise stain fades away
CONTROLS
6.
HCl extraction: Treat control section with 0.2 M acetate buffer at pH 4.2. (Mix 0.88
ml acetic acid 0.44 g of anhydrous sodium acetate in 100 ml, adjust pH to 4.2). Then
immerse section in 1N HCl at 60 0C for exactly 5 min to selectively extract RNA.
(This extraction of RNA is not recommended). Then proceeds with staining.
7.
Perchloric acid extraction: Treatment in 1N perchloric acid (or make 10.94 % v/v
commercially available 60 % perchloric acid) at 4 oC for 12-24 hrs removes RNA
selectively. A treatment in 0.5N perchloric acid at 70 oC for 20-40 min. removes total
nuclei acid. After each or both of these extractions, place the section in 1 % v/v
sodium carbonate for 5 min, wash in running water and then proceeds with staining.
Try the other methods
AZUR B Method: This method is meant for both DNA and RNA. Hydrate the section fixed in
Carnoy’s fluid or Acetic alcohol. Place the section in 0.25 mg /ml solution of Azur B in
citrate buffer at pH 4.0 for1- 2 hrs at 50 oC. Wash in water and place in pure tertiary butanol
for 30 min. Give two or more changes in butanol. Leave overnight if stained if stained too
deep in butanol. DNA containing structures appear greenish blue while RNA sites appear
purple or dark blue. To prepare Citrate buffer (pH 4) add 33 ml of stock A with 17 ml of
stock B and made up the volume to 100 ml with distilled water. (Stock A-0.1 M Soln. of
Citrate Acid i.e. 21.01 g in 1000 ml; Stock B 0.1 M Soln. of Sodium Citrate -29.41 g C6H5O7
Na3. 2H2O in 1000 ml). Azur B also stain lignin greenish blue). Cuticle and sieve plates
often appear red.
Gallocyanin-Chrome Alum Method: This method is meant for total nucleic acids. Stain freshly
prepared callocyanin-chrome alum solution for 10 min and wash in water for 30 min.
Dehydrate and then pass through xylene before mounting. Nuclei acid stains bluish violet. To
prepare the stain dissolve 5 g of chrome alum in 100 ml water. Add 150 mg Gallocyanin and
boil the solution. After filter, make up the volume to 100 ml and adjust the ph to 1.64.
Pyronin Y method : This is for RNA. Hydrate the section of the fixed materials in acetic acid
ethanol or neutral buffered Formaldehyde. Stain in 2 % aqueous Pyronin Y solution for 2
min-wash in water-blot the back and uncovered areas of the slide-give three rapid dips in
absolute ethanol 1 and then in absolute ethanol 2 for 30 s. Immerse for 30 s in 1:1 absolute
ethanol:xylene. Pass through xylene and mount in DPX. RNA stains Pink. To prepare 2 %
Pyronin Y stain dissolve 1 g stain in 50 ml water. Extract the solution 5 times with
Chloroform for removing impurities. For this agitate the stain with equal volume of
Chloroform in a separating funnel, let it stand and discard the heavier layer (lower layer) of
chloroform. Repeat the process for 5 times till chloroform layer remains somewhat
colourless. Leave the stain overnight in a open beaker to evaporate the left over chloroform.
Next day replenish the Pyronin Y to 50 ml by adding water. Use this staining mixture fresh.
95
Suggested Readings
1. Baker, J.R. 1956. The histochemical recognistion of phenols, especially tyrosine.
Quart. J. Microscop. Sci. 97:161
2. Bendre, A. and Kumar, A. 1975. A Text Book of Practical Botany. Vol II. Rastogi
Publication, Meerut, 384 p.
3. Bhatnagar, S.S. 1976. A Class Book of Practical Botany. Vol- 2. Fourth edition,
Ratan Prakasan Mandir, Agra, 546 p
4. Burstone, N.S. 1962. Enzyme Histochemistry and its Application to the Study of
Neoplasm. Academic Press, New York and London
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