ARRANGEMENT OF VASCULAR BUNDLES
Types of fibro-vascular bundles 292
Radial vascular bundles 292
Concentric vascular bundles 295
Collateral vascular bundles 295
Bi-collateral vascular bundles 298
Open collateral vascular bundles 298
INDEX
TABLE OF ILLUSTRATIONS
PAGE
FIG. 1. Tripod Magnifier 4
FIG. 2. Watchmaker’s Loupe 4
FIG. 3. Folding Magnifier 4
FIG. 4. Reading Glass 4
FIG. 5. Steinheil Aplanatic Lens 5
FIG. 6. Dissecting Microscope 5
FIG. 7. Compound Microscope of Robert Hooke 8
FIG. 8. Compound Microscope 10
FIG. 9. Abbé Condenser 11
FIG. 10. 11
FIG. 11. 11
FIG. 12. Objectives 11
FIG. 13. 12
FIG. 14. 12
FIG. 15. Eye-Pieces. 12
FIG. 16. Pharmacognostic Microscope 12
FIG. 17. Research Microscope 14
FIG. 18. Special Research Microscope 14
FIG. 19. Greenough Binocular Microscope 15
FIG. 20. Polarization Microscope 16
FIG. 21. Ocular Micrometer 19
FIG. 22. Stage Micrometer 19
FIG. 23. Micrometer Eye-Piece 20
FIG. 24. Micrometer Eye-Piece 21
FIG. 25. Mechanical Stage 22
FIG. 26. Camera Lucida 22
FIG. 27. Camera Lucida 22
FIG. 28. Drawing Apparatus 23
FIG. 29. Microphotographic Apparatus 24
FIG. 30. Micro Lamp 27
FIG. 31. Paraffin-embedding Oven 30
FIG. 32. Paraffin Blocks 31
FIG. 33. Hand Microtome 31
FIG. 34. Hand Cylinder Microtome 34
FIG. 35. Hand Table Microtome 34
FIG. 36. Base Sledge Microtome 35
FIG. 37. Minot Rotary Microtome 36
FIG. 38. Reagent Set 39
FIG. 39. Measuring Cylinder 40
FIG. 40. Staining Dish 40
FIG. 41. Round Cover Glass 44
FIG. 42. Square Cover Glass 44
FIG. 43. Rectangular Cover Glass 44
FIG. 44. Glass Slide 44
FIG. 45. Histological Forceps 45
FIG. 46. Forceps 45
FIG. 47. Sliding-pin Forceps 45
FIG. 48. Dissecting Needle 46
FIG. 49. Scissors 46
FIG. 50. Scalpels 47
FIG. 51. Turntable 47
FIG. 52. Slide Box 48
FIG. 53. Slide Tray 48
FIG. 54. Slide Cabinet 49
PLATE 1 THE ONION ROOT 56
PLATE 2 LEAF EPIDERMIS 60
PLATE 3 LEAF EPIDERMIS 61
PLATE 4 TESTA EPIDERMAL CELLS 64
PLATE 5 TESTA CELLS 65
PLATE 6 PAPILLÆ 68
PLATE 7 UNICELLULAR SOLITARY HAIRS 70
PLATE 8 CLUSTERED UNICELLULAR HAIRS 71
PLATE 9 MULTICELLULAR UNISERIATE NON-BRANCHED HAIRS 73
PLATE 10 MULTICELLULAR MULTISERIATE NON-BRANCHED HAIRS 75
PLATE 11 MULTICELLULAR UNISERIATE BRANCHED HAIRS 76
PLATE 12 NON-GLANDULAR MULTICELLULAR HAIRS 78
PLATE 13 MULTICELLULAR MULTISERIATE BRANCHED HAIRS 79
PLATE 14 MULTICELLULAR MULTISERIATE BRANCHED HAIRS 81
PLATE 15 MULTICELLULAR MULTISERIATE BRANCHED HAIRS 82
PLATE 16 PERIDERM OF CASCARA SAGRADA (_Rhamnus purshiana_,
D.C.) 84
PLATE 17 MANDRAKE RHIZOME and WHITE CINNAMON 86
PLATE 18 PERIDERM OF WHITE OAK (_Quercus alba_, L.) 87
PLATE 19 CRYSTAL-BEARING FIBRES OF BARKS 91
PLATE 20 CRYSTAL-BEARING FIBRES OF BARKS 93
PLATE 21 CRYSTAL-BEARING FIBRES OF LEAVES 94
PLATE 22 BRANCHED BAST FIBRES 95
PLATE 23 POROUS AND STRIATED BAST FIBRES 97
PLATE 24 POROUS AND NON-STRIATED BAST FIBRES 98
PLATE 25 NON-POROUS AND STRIATED BAST FIBRES 99
PLATE 26 NON-POROUS AND NON-STRIATED BAST FIBRES 101
PLATE 27 GROUPS OF BAST FIBRES 102
PLATE 28 WOOD FIBRES 105
PLATE 29 CATNIP STEM and MOTHERWORT STEM 107
PLATE 30 COLLENCHYMA CELLS 108
PLATE 31 BRANCHED STONE CELLS 110
PLATE 32 POROUS AND STRIATED STONE CELLS 113
PLATE 33 POROUS AND NON-STRIATED STONE CELLS 114
PLATE 34 CINNAMON, RUELLA ROOT, CASCARA and CINNAMON 115
PLATE 35 CROSS-SECTIONS OF ENDODERMAL CELLS OF 117
PLATE 36 LONGITUDINAL SECTIONS OF ENDODERMAL CELLS 119
PLATE 37 HYPODERMAL CELLS 120
PLATE 38 CROSS-SECTION OF SARSAPARILLA ROOT (_Smilax
officinalis_, Kunth) 123
PLATE 39 ROOT HAIRS (Fragments) 124
PLATE 40 ANNULAR AND SPIRAL VESSELS 129
PLATE 41 SPIRAL VESSELS 130
PLATE 42 SCLARIFORM VESSELS 132
PLATE 43 RETICULATE VESSELS 133
PLATE 44 PITTED VESSELS 134
PLATE 45 VESSELS 135
PLATE 46 SIEVE TUBE 137
PLATE 47 RADIAL LONGITUDINAL SECTION OF WHITE SANDALWOOD
(_Santalum album_, L.) 140
PLATE 48 KAVA-KAVA ROOT and WHITE PINE BARK 143
PLATE 49 BLACK INDIAN HEMP and BLACK INDIAN HEMP ROOT 145
PLATE 50 LATEX VESSELS 146
PLATE 51 PARENCHYMA CELLS 148
PLATE 52 GRINDELIA STEM (longitudinal) and GRINDELIA STEM
(cross-section) 149
PLATE 53 ACONITE STEM and PEPPERMINT STEM 152
PLATE 54 TYPES OF STOMA 153
PLATE 55 LEAF EPIDERMI WITH STOMA 155
PLATE 56 BELLADONNA LEAF, DEER TONGUE LEAF and WHITE PINE LEAF 156
PLATE 57 ELDER BARK 159
PLATE 58 INTERCELLULAR AIR SPACES 160
PLATE 59 IRREGULAR INTERCELLULAR AIR SPACES 161
PLATE 60 GLANDULAR HAIRS 165
PLATE 61 STALKED GLANDULAR HAIRS 167
PLATE 62 CALAMUS RHIZOME and WHITE PINE BARK 169
PLATE 63 CANELLA ALBA BARK and KLIP BUCHU LEAF 170
PLATE 64 BITTER ORANCE PEEL and WHITE PINE LEAF 171
PLATE 65 CINNAMON, CALUMBA, PARENCHYMA, SARSAPARILLA,
LEPTANDRA, QUEBRACHO, BLACKBERRY 174
PLATE 66 MUCILAGE AND RESIN 175
PLATE 67 CROSS-SECTION OF SKUNK-CABBAGE LEAF (_Symplocarpus
fœtidus_, [L.] Nutt.) 177
PLATE 68 RESERVE CELLULOSE 180
PLATE 69 RESERVE CELLULOSE 181
PLATE 70 STARCH 186
PLATE 71 STARCH 187
PLATE 72 STARCH 189
PLATE 73 STARCH 190
PLATE 74 STARCH 191
PLATE 75 STARCH GRAINS 192
PLATE 76 STARCH MASSES 193
PLATE 77 INULIN (_Inula helenium_, L.) 195
PLATE 77_a_ ALEURONE GRAINS 199
PLATE 78 MICRO-CRYSTALS 201
PLATE 79 RAPHIDES 203
PLATE 80 ROSETTE CRYSTALS 204
PLATE 81 INCLOSED ROSETTE CRYSTALS 206
PLATE 82 SOLITARY CRYSTAL 207
PLATE 83 SOLITARY CRYSTALS 208
PLATE 84 SOLITARY CRYSTALS 209
PLATE 85 SOLITARY CRYSTALS 211
PLATE 86 SOLITARY CRYSTALS 212
PLATE 87 ROSETTE CRYSTALS AND SOLITARY CRYSTALS OCCURRING IN 213
PLATE 88 CYSTOLITHS 214
PLATE 89 CROSS-SECTION OF ROOT OF SPIGELIA MARYLANDICA, L. 220
PLATE 90 RUELLIA ROOT (_Ruellia ciliosa_, Pursh.). 222
PLATE 91 CROSS-SECTION OF RHIZOME OF SPIGELIA MARYLANDICA, L. 224
PLATE 92 CROSS-SECTION OF RHIZOME OF RUELLIA CILIOSA, Pursh. 225
PLATE 93 POWDERED SPIGELIA MARYLANDICA, L. 228
PLATE 94 POWDERED RUELLIA CILIOSA, Pursh. 229
PLATE 95 CROSS-SECTION OF STEM OF SPIGELIA MARYLANDICA, L. 234
PLATE 96 CROSS-SECTION OF STEM OF RUELLIA CILIOSA, Pursh. 236
PLATE 97 POWDERED HOREHOUND (_Marrubium vulgare_, L). 238
PLATE 98 SPURIOUS HOREHOUND (_Marrubium peregrinum_, L.) 239
PLATE 99 POWDERED INSECT FLOWER STEMS (_Chrysanthemum
cinerariifolium_, [Trev.], Vis.) 240
PLATE 100 CROSS-SECTION OF BUCHU STEMS (_Barosma betulina_
[Berg.], Barth, and Wendl.) 243
PLATE 101 BUCHU STEM and LEPTANDRA RHIZOME 244
PLATE 102 POWDERED BUCHU STEMS (_Barosma betulina_ [Berg.],
Barth. and Wendl.). 246
PLATE 103 CROSS-SECTION OF UNROSSED WHITE PINE BARK (_Pinus
strobus_, L.) 249
PLATE 104 POWDERED WHITE PINE BARK (_Pinus strobus_, L.) 251
PLATE 105 CROSS-SECTION OF QUASSIA WOOD (_Picræna excelsa_
[Sw.], Lindl.) 255
PLATE 106 TANGENTIAL SECTION OF QUASSIA WOOD (_Picræna
excelsa_ [Sw.], Lindl.) 256
PLATE 107 RADIAL SECTION OF QUASSIA WOOD (_Picræna excelsa_
[Sw.], Lindl.) 257
PLATE 108 CROSS-SECTION OF KLIP BUCHU JUST OVER THE VEIN 261
PLATE 109 POWDERED KLIP BUCHU 263
PLATE 110 CROSS-SECTION MOUNTAIN LAUREL (_Kalmia latifolia_,
L.) 265
PLATE 111 CROSS-SECTION TRAILING ARBUTUS LEAF (_Epigæa
repens_, L.) 266
PLATE 112 POWDERED INSECT FLOWER LEAVES 268
PLATE 113 SMOOTH-WALLED POLLEN GRAINS 271
PLATE 114 SPINY WALLED POLLEN GRAINS 272
PLATE 115 PAPILLÆ 275
PLATE 116 PAPILLÆ OF STIGMAS 276
PLATE 117 PAPILLÆ OF STIGMAS 277
PLATE 118 POWDERED CLOSED INSECT FLOWER 279
PLATE 119 POWDERED OPEN INSECT FLOWER 281
PLATE 120 POWDERED WHITE DAISIES (_Chrysanthemum
leucanthemum_, L.) 283
PLATE 121 CROSS-SECTION OF CELERY FRUIT (_Apium
graveolens_, L.) 286
PLATE 121 CROSS-SECTION OF CELERY FRUIT (_Apium
graveolens_, L.) 286
PLATE 123 CROSS-SECTION SWEET ALMOND SEED 290
PLATE 124 CROSS-SECTION OF A RADIAL VASCULAR BUNDLE OF
SKUNK CABBAGE ROOT 293
PLATE 125 CROSS-SECTION OF A PHLOEM-CENTRIC BUNDLE OF
CALAMUS RHIZOME (_Acorus calamus_, L.) 294
PLATE 126 CROSS-SECTION OF A CLOSED COLLATERAL BUNDLE OF
MANDRAKE STEM (_Podophyllum peltatum_, L.) 286
PLATE 127 BI-COLLATERAL BUNDLE OF PUMPKIN STEM (_Curcurbita
pepo_, L.) 297
Part I
SIMPLE AND COMPOUND MICROSCOPES AND MICROSCOPIC TECHNIC
CHAPTER I
THE SIMPLE MICROSCOPES
The construction and use of the =simple microscope= (magnifiers)
undoubtedly date back to very early times. There is sufficient
evidence to prove that spheres of glass were used as burning spheres
and as magnifiers by people antedating the Greeks and Romans.
The simple microscopes of to-day have a very wide range of
application and a corresponding variation in structure and in
appearance.
Simple microscopes are used daily in classifying and studying crude
drugs, testing linen and other cloth, repairing watches, in reading,
and identifying insects. The more complex simple microscopes are used
in the dissection and classification of flowers.
The =watchmaker’s loupe=, the =linen tester=, the =reading glass=,
the =engraver’s lens=, and the simplest folding magnifiers consist
of a double convex lens. Such a lens produces an erect, enlarged
image of the object viewed when the lens is placed so that the object
is within its focal distance. The focal distance of a lens varies
according to the curvature of the lens. The greater the curvature,
the shorter the focal distance and the greater the magnification.
The more complicated simple microscope consists of two or more
lenses. The double and triple magnifiers consist of two and three
lenses respectively.
When an object is viewed through three lenses, the magnification is
greater than when viewed through one or two lenses, but a smaller
part of the object is magnified.
FORMS OF SIMPLE MICROSCOPES
TRIPOD MAGNIFIER
The =tripod magnifier= (Fig. 1) is a simple lens mounted on a
mechanical stand. The tripod is placed over the object and the focus
is obtained by means of a screw which raises or lowers the lens,
according to the degree it is magnified.
WATCHMAKER’S LOUPE
The =watchmaker’s loupe= (Fig. 2) is a one-lens magnifier mounted on
an ebony or metallic tapering rim, which can be placed over the eye
and held in position by frowning or contracting the eyelid.
[Illustration: FIG. 1.--Tripod Magnifier]
[Illustration: FIG. 2.--Watchmaker’s Loupe]
FOLDING MAGNIFIER
The =folding magnifier= (Fig. 3) of one or more lenses is mounted
in such a way that, when not in use, the lenses fold up like the
blade of a knife, and when so folded are effectively protected from
abrasion by the upper and lower surfaces of the folder.
[Illustration: FIG. 3.--Folding Magnifier]
[Illustration: FIG. 4.--Reading Glass]
READING GLASSES
=Reading glasses= (Fig. 4) are large simple magnifiers, often six
inches in diameter. The lens is encircled with a metal band and
provided with a handle.
[Illustration: FIG. 5.--Steinheil Aplanatic Lens]
STEINHEIL APLANATIC LENSES
=Steinheil aplanatic lenses= (Fig. 5) consist of three or four
lenses cemented together. The combination is such that the field is
large, flat, and achromatic. These lenses are suitable for field,
dissecting, and pocket use. When such lenses are placed in simple
holders, they make good dissecting microscopes.
[Illustration: FIG. 6.--Dissecting Microscope]
DISSECTING MICROSCOPE
The =dissecting microscope= (Fig. 6) consists of a Steinheil lens
and an elaborate stand, a firm base, a pillar, a rack and pinion,
a glass stage, beneath which there is a groove for holding a metal
plate with one black and one white surface. The nature of the object
under observation determines whether a plate is used. When the plate
is used and when the object is studied by reflected light it is
sometimes desirable to use the black and sometimes the white surface.
The mirror, which has a concave and a plain surface, is used to
reflect the light on the glass stage when the object is studied by
transmitted light. The dissecting microscope magnifies objects up to
twenty diameters, or twenty times their real size.
CHAPTER II
COMPOUND MICROSCOPES
The =compound microscope= has undergone wonderful changes since 1667,
the days of Robert Hooke. When we consider the crude construction
and the limitations of Robert Hooke’s microscope, we marvel at the
structural perfection and the unlimited possibilities of the modern
instrument. The advancement made in most sciences has followed the
gradual perfection of this instrument.
The illustration of Robert Hooke’s microscope (Fig. 7) will convey
to the mind more eloquently than words the crudeness of the early
microscopes, especially when it is compared with the present-day
microscopes.
STRUCTURE OF THE COMPOUND MICROSCOPE
The parts of the compound microscope (Fig. 8) may be grouped
into--first, the mechanical, and, secondly, into the optical parts.
THE MECHANICAL PARTS
1. The =foot= is the basal part, the part which supports all the
other mechanical and optical parts. The foot should be heavy enough
to balance the other parts when they are inclined. Most modern
instruments have a three-parted or tripod-shaped base.
2. The =pillar= is the vertical part of the microscope attached to
the base. The pillar is joined to the limb by a hinged joint. The
hinges make it possible to incline the microscope at any angle, thus
lowering its height. In this way, short, medium, and tall persons
can use the microscope with facility. The part of the pillar above
the hinge is called the _limb_. The limb may be either straight
or curved. The curved form is preferable, since it offers a more
suitable surface to grasp in transferring from box or shelf to the
desk, and _vice versa_.
[Illustration: FIG. 7.--Compound Microscope of Robert Hooke]
3. The =stage= is either stationary or movable, round or square, and
is attached to the limb just above the hinge. The upper surface is
made of a composition which is not easily attacked by moisture and
reagents. The centre of the stage is perforated by a circular opening.
4. The =sub-stage= is attached below the stage and is for the purpose
of holding the iris diaphragm and Abbé condenser. The raising and
lowering of the sub-stage are accomplished by a rack and pinion.
5. The =iris diaphragm=, which is held in the sub-stage below the
Abbé condenser, consists of a series of metal plates, so arranged
that the light entering the microscope may be cut off completely or
its amount regulated by moving a control pin.
6. The =fine adjustment= is located either at the side or at the top
of the limb. It consists of a fine rack and pinion, and is used in
focusing an object when the low-power objective is in position, or in
finding and focusing the object when the high-power objective is in
position.
7. The =coarse adjustment= is a rack and pinion used in raising and
lowering the body-tube and in finding the approximate focus when
either the high- or low-power objective is in position.
8. The =body-tube= is the path traveled by the rays of light entering
the objectives and leaving by the eye-piece. To the lower part of the
tube is attached the nose-piece, and resting in its upper part is the
draw-tube, which holds the eye-piece. On the outer surface of the
draw-tube there is a scale which indicates the distance it is drawn
from the body-tube.
9. The =nose-piece= may be simple, double, or triple, and it is
protected from dust by a circular piece of metal. Double and triple
nose-pieces may be revolved, and like the simple nose-piece they hold
the objectives in position.
THE OPTICAL PARTS
1. The =mirror= is a sub-stage attachment one surface of which is
plain and the other concave. The plain surface is used with an Abbé
condenser when the source of light is distant, while the concave
surface is used with instruments without an Abbé condenser when the
source of light is near at hand.
[Illustration: FIG. 8.--Compound Microscope
Eyepiece
Draw Tube
Body Tube
Coarse
Adjustment
Revolving Nosepiece for three Objectives
Fine Adjustment
Stage
Objectives
Limb
Abbi Condenser
Iris Diaphragm
Hinge for Inclining
Substage Attachment
Mirror
Pillar
Foot]
2. The =Abbé condenser= (Fig. 9) is a combination of two or more
lenses, arranged so as to concentrate the light on the specimen
placed on the stage. The condenser is located in the opening of the
stage, and its uppermost surface is circular and flat.
[Illustration: FIG. 9--Abbé Condenser]
3. =Objectives= (Figs. 10, 11, and 12). There are low, medium, and
high-power objectives. The low-power objectives have fewer and
larger lenses, and they magnify least, but they show more of the
object than do the high-power objectives. There are three chief
types of objectives: First, dry objectives; second, wet objectives,
of which there are the water-immersion objectives; and third, the
oil-immersion objectives. The dry objectives are used for most
histological and pharmacognostical work. For studying smaller objects
the water objective is sometimes desirable, but in bacteriological
work the oil-immersion objective is almost exclusively used. The
globule of water or oil, as the case may be, increases the amount of
light entering the objective, because the oil and water bend many
rays into the objective which would otherwise escape.
[Illustration: FIG. 10.]
[Illustration: FIG. 11.]
[Illustration: FIG. 12. Objectives.]
4. =Eye-pieces= (Figs. 13, 14, and 15) are of variable length, but
structurally they are somewhat similar. The eye-piece consists of a
metal tube with a blackened inner tube. In the centre of this tube
there is a small diaphragm for holding the ocular micrometer. In the
lower end of the tube a lens is fastened by means of a screw. This,
the field lens, is the larger lens of the ocular. The upper, smaller
lens is fastened in the tube by a screw, but there is a projecting
collar which rests, when in position, on the draw-tube.
[Illustration: FIG. 13.]
[Illustration: FIG. 14.]
[Illustration: FIG. 15. Eye-Pieces.]
The longer the tube the lower the magnification. For instance, a
two-inch ocular magnifies less than an inch and a half, a one-inch
less than a three-fourths of an inch, etc.
The greater the curvature of the lenses of the ocular the higher will
be the magnification and the shorter the tube-length.
FORMS OF COMPOUND MICROSCOPES
The following descriptions refer to three different models of
compound microscopes: one which is used chiefly as a pharmacognostic
microscope, one as a research microscope stand, while the third type
represents a research microscope stand of highest order, which is
used at the same time for taking microphotographs.
[Illustration: FIG. 16.--Pharmacognostic Microscope]
PHARMACOGNOSTIC MICROSCOPE
The =pharmacognostic microscope= (Fig. 16) is an instrument
which embodies only those parts which are most essential for the
examination of powdered drugs, bacteria, and urinary sediments. This
microscope is provided with a stage of the dimensions 105 × 105 mm.
This factor and the distance of 80 mm. from the optical centre to the
handle arm render it available for the examination of even very large
objects and preparations, or preparations suspended in glass dishes.
The stand is furnished with a side micrometer, a fine adjustment
having knobs on both sides, thereby permitting the manipulation of
the micrometer screw either by left or right hand. The illuminating
apparatus consists of the Abbé condenser of numerical aperture
of 1.20, to which is attached an iris diaphragm for the proper
adjustment of the light. A worm screw, mounted in connection with the
condenser, serves for the raising and lowering of the condenser, so
that the cone of illuminating pencils can be arranged in accordance
to the objective employed and to the preparation under observation.
The objectives necessary are those of the achromatic type, possessing
a focal length of 16.2 mm. and 3 mm. Oculars which render the best
results in regard to magnification in connection with the two
objectives mentioned are the Huyghenian eye-pieces II and IV so that
magnifications are obtained varying from 62 to 625. It is advisable,
however, to have the microscope equipped with a triple revolving
nose-piece for the objectives, so that provision is made for the
addition of an oil-immersion objective at any time later should the
microscope become available for bacteriological investigations.
THE RESEARCH MICROSCOPE
The =research microscope= used in research work (Fig. 17) must be
equipped more elaborately than the microscope especially designed for
the use of the pharmacognosist. While the simple form of microscope
is supplied with the small type of Abbé condenser, the research
microscope is furnished with a large illuminating apparatus of
which the iris diaphragm is mounted on a rack and pinion, allowing
displacement obliquely to the optical centre, also to increase
resolving power in the objectives when observing those objects which
cannot be revealed to the best advantage with central illumination.
Another iris is furnished above the condenser; this iris becomes
available the instant an object is to be observed without the aid
of the condenser, in which case the upper iris diaphragm allows
proper adjustment of the light. The mirror, one side plane, the
other concave, is mounted on a movable bar, along which it can be
slid--another convenience for the adjustment of the light. The
microscope stage of this stand is of the round, rotating and centring
pattern, which permits a limited motion to the object slide: The
rotation of the microscope stage furnishes another convenience in the
examination of objects in polarized light, allowing the preparation
to be rotated in order to distinguish the polarization properties of
the objects under observation.
[Illustration: FIG. 17.--Research Microscope]
[Illustration: FIG. 18.--Special Research Microscope]
SPECIAL RESEARCH MICROSCOPE
A =special research microscope= of the highest order (Fig. 18)
is supplied with an extra large body tube, which renders it of
special advantage for micro-photography. Otherwise in its mechanical
equipment it resembles very closely the medium-sized research
microscope stand, with the exception that the stand is larger in its
design, therefore offering universal application. In regard to the
illuminating apparatus, it is advisable to mention that the one in
the large research microscope stand is furnished with a three-lens
condenser of a numerical aperture of 1.40, while the medium-sized
research stand is provided with a two-lens condenser of a numerical
aperture of 1.20. The stage of the microscope is provided with a
cross motion--the backward and forward motion of the preparation is
secured by rack and pinion, while the side motion is controlled by a
micrometric worm screw. In cases where large preparations are to be
photographed, the draw-tube with ocular and the slider in which the
draw-tubes glide are removed to allow the full aperture of wide-angle
objectives to be made use of.
[Illustration: FIG. 19.--Greenough Binocular Microscope]
BINOCULAR MICROSCOPE
The =Greenough binocular microscope=, as shown in Fig. 19, consists
of a microscope stage with two tubes mounted side by side and moving
on the same rack and pinion for the focusing adjustment. Either tube
can be used without the other. The oculars are capable of more or
less separation to suit the eyes of different observers. In each
of the drub-like mountings, near the point where the oculars are
introduced, porro-prisms have been placed, which erect the image.
This microscope gives most perfect stereoscopic images, which are
erect instead of inverted, as in the monocular compound microscopes.
The Greenough binocular microscope is especially adapted for
dissection and for studying objects of considerable thickness.
POLARIZATION MICROSCOPE
The =polarization microscope= (Fig. 20) is used chiefly for the
examination of crystals and mineral sections as well as for the
observation of organic bodies in polarized light. It can, however,
also be used for the examination of regular biological preparations.
[Illustration: FIG. 20.--Polarization Microscope]
If compared with the regular biological microscope, the polarization
microscope is found characteristic of the following points: it is
supplied with a polarization arrangement. The latter consists of a
polarizer and analyzer. The polarizer is situated in a rotating mount
beneath the condensing system. The microscope, of which the diagram
is shown, possesses a triple “Ahrens” prism of calcite. The entering
light is divided into two polarized parts, situated perpendicularly
to each other. The so-called “ordinary” rays are reflected to one
side by total reflection, which takes place on the inner cemented
surface of the triple prism, allowing the so-called “extraordinary”
rays to pass through the condenser. If the prism is adjusted to
its focal point, it is so situated that the vibration plane of the
extra-ordinary rays are in the same position as shown in the diagram
of the illustration.
The analyzer is mounted within the microscope-tube above the
objective. Situated on a sliding plate, it can be shifted into
the optical axis whenever necessary. The analyzer consists of a
polarization prism after Glan-Thompson. The polarization plane of the
active extraordinary rays is situated perpendicularly to the plane as
shown in the diagram. The polarization prisms are ordinarily crossed.
In this position the field of the microscope is darkened as long as
no substance of a double refractive index has been introduced between
the analyzer and polarizer. In rotating the polarizer up to the mark
90, the polarization prisms are mounted parallel and the field of
the microscope is lighted again. Immediately above the analyzer and
attached to the mounting of the analyzer a lens of a comparatively
long focal length has been placed in order to overcome the difference
in focus created by the introduction of the analyzer into the optical
rays.
The condensing system is mounted on a slider, and, furthermore,
can be raised and lowered along the optical centre by means of a
rack-and-pinion adjustment. If lowered sufficiently, the condensing
system can be thrown to the side to be removed from the optical
rays. The condenser consists of three lenses. The two upper lenses
are separately mounted to an arm, which permits them to be tilted to
one side in order to be removed from the optical rays. The complete
condenser is used only in connection with high-power objectives.
As far as low-power objectives are concerned, the lower condensing
lens alone is made use of, and the latter is found mounted to the
polarizer sleeve. Below the polarizer and above the lower condensing
lens an iris diaphragm is found.
The microscope table is graduated on its periphery, and, furthermore,
carries a vernier for more exact reading.
The polarization microscope is not furnished with an objective
nose-piece. Every objective, however, is supplied with an individual
centring head, which permits the objective to be attached to
an objective clutch-changer, situated at the lower end of the
microscope-tube. The centring head permits the objectives to be
perfectly centred and to remain centred even if another objective is
introduced into the objective clutch-changer.
At an angle of 45 degrees to the polarization plane of polarizer and
analyzer, a slot has been provided, which serves for the introduction
of compensators.
Between analyzer and ocular, another slot is found which permits
the Amici-Bertrand lens to be introduced into the optical axis. The
slider for the Bertrand lens is supplied with two centring screws
whereby this lens can be perfectly and easily centred. The Bertrand
lens serves the purpose of observing the back focal plane of the
microscope objective. In order to allow the Bertrand lens to be
focused, the tube can be raised and lowered for this purpose. An iris
diaphragm is mounted above the Bertrand lens.
If the Bertrand lens is shifted out of the optical axis, one can
observe the preparation placed upon the microscope stage and,
depending on its thickness or its double refraction, the interference
color of the specimen. This interference figure is called the
orthoscopic image and, accordingly, one speaks of the microscope as
being used as an “orthoscope.”
After the Bertrand lens has been introduced into the optical axis,
the interference figure is visible in the back focal plane of the
objective. Each point of this interference figure corresponds to
a certain direction of the rays of the preparation itself. This
arrangement permits observation of the change of the reflection of
light taking place in the preparation, this in accordance with the
change of the direction of the rays. This interference figure is
called the conoscopic image, and, accordingly, the microscope is used
as a “conoscope.”
Many types of polarization microscopes have been constructed; those
of a more elaborate form are used for research investigations; others
of smaller design for routine investigations.
CHAPTER III
MICROSCOPIC MEASUREMENTS
In making critical examinations of powdered drugs, it is frequently
necessary to measure the elements under observation, particularly in
the case of starches and crystals.
[Illustration: FIG. 21.--Ocular Micrometer]
OCULAR MICROMETER
Microscopic measurements are made by the =ocular micrometer= (Fig.
21). This consists of a circular piece of transparent glass on the
centre of which is etched a one- or two-millimeter scale divided into
one hundred or two hundred divisions respectively. The value of each
line is determined by standardizing with a stage micrometer.
STAGE MICROMETER
[Illustration: FIG. 22.--Stage Micrometer]
The =stage micrometer= (Fig. 22) consists of a glass slide upon which
is etched a millimeter scale divided into one hundred equal parts or
lines: each line has a value of one hundredth of a millimeter.
STANDARDIZATION OF OCULAR MICROMETER WITH LOW-POWER OBJECTIVE
Having placed the ocular micrometer in the eye-piece and the stage
micrometer on the centre of the stage, focus until the lines of the
stage micrometer are clearly seen. Then adjust the scales until the
lines of the stage micrometer are parallel with and directly under
the lines of the ocular micrometer.
Ascertain the number of lines of the stage micrometer covered by the
one hundred lines of the ocular micrometer. Then calculate the value
of each line of the ocular. This is done in the following manner:
If the one hundred lines of the ocular cover seventy-five lines
of the stage micrometer, then the one hundred lines of the ocular
micrometer are equivalent to seventy-five one-hundredths, or
three-fourths, of a millimeter. One line of the ocular micrometer
will therefore be equivalent to one-hundredth of seventy-five
one-hundredths, or .0075 part of a millimeter, and as a micron is the
unit for measuring microscopic objects, this being equivalent to one
one-thousandth of a millimeter, the value of each line of the ocular
will therefore be 7.5 microns.
[Illustration: FIG. 23.--Micrometer Eye-Piece]
With the high-power objective in place, ascertain the value of each
line of the ocular. If one hundred lines of the ocular cover only
twelve lines of the stage micrometer, then the one hundred lines of
the ocular are equivalent to twelve one-hundredths of a millimeter,
the value of one line being equivalent to one one-hundredth of twelve
one-hundredths, or twelve ten-thousandths of a millimeter, or .0012,
or 1.2_µ_.
It will therefore be seen that objects as small as a thousandth of a
millimeter can be accurately measured by the ocular micrometer.
In making microscopic measurements it is only necessary to find how
many lines of the ocular scale are covered by the object. The number
of lines multiplied by the equivalent of each line will be the size
of the object in microns, or _micromillimeters_.
[Illustration: FIG. 24.--Micrometer Eye-Piece]
MICROMETER EYE-PIECES
=Micrometer eye-pieces= (Figs. 23 and 24) may be used in making
measurements. These eye-pieces with micrometer combinations are
preferred by some workers, but the ocular micrometer will meet the
needs of the average worker.
MECHANICAL STAGES
Moving objects by hand is tiresome and unsatisfactory, first, because
of the possibility of losing sight of the object under observation,
and secondly, because the field cannot be covered so systematically
as when a mechanical stage is used for moving slides.
The =mechanical stage= (Fig. 25) is fastened to the stage by a screw.
The slide is held by two clamps. There is a rack and pinion for
moving the slide to left or right, and another rack and pinion for
moving the slide forward and backward.
[Illustration: FIG. 25. Mechanical Stage]
CAMERA LUCIDA
The =camera lucida= is an optical mechanical device for aiding the
worker in making drawings of microscopic objects. The instrument is
particularly necessary in research work where it is desirable to
reproduce an object in all its details. In fact, all reproductions
illustrating original work should be made by means of the camera
lucida or by microphotography.
[Illustration: FIG. 26.--Camera Lucida]
[Illustration: FIG. 27.--Camera Lucida]
A great many different types of camera lucidas or drawing apparatus
are obtainable, varying from simple-inexpensive to complex-expensive
forms. Figs. 26, 27, and 28 show simple and complex forms.
[Illustration: FIG. 28.--Drawing Apparatus]
MICROPHOTOGRAPHIC APPARATUS
The =microphotographic apparatus= (Fig. 29), as the name implies, is
an apparatus constructed in such a manner that it may be attached to
a microscope when we desire to photograph microscopic objects. It
consists of a metal base and a polished metal pillar for holding the
bellows, slide holder, ground-glass observation plate, and eye-piece.
In making photographs, the small end of the bellows is attached to
the ocular of the microscope, the focus adjusted, and the object or
objects photographed. More uniform results are obtained in making
such photographs if an artificial light of an unvarying candle-power
is used.
[Illustration: FIG. 29.--Microphotographic Apparatus]
There are obtainable more elaborate microphotographic apparatus than
the one figured and described, but for most workers this one will
prove highly satisfactory. It is possible, by inclining the tube of
the microscope, to make good microphotographs with an ordinary plate
camera. This is accomplished by removing the lens of the camera and
attaching the bellows to the ocular, focusing, and photographing.
CHAPTER IV
HOW TO USE THE MICROSCOPE
In beginning work with the compound microscope, place the base of the
microscope opposite your right shoulder, if you are right-handed; or
opposite your left shoulder, if you are left-handed. Incline the body
so that the ocular is on a level with your eye, if necessary; but
if not, work with the body of the microscope in an erect position.
In viewing the specimen, keep both eyes open. Use one eye for
observation and the other for sketching. In this way it will not be
necessary to remove the observation eye from the ocular unless it be
to complete the details of a sketch.
=Learn to use both eyes.= Most workers, however, accustom themselves
to using one eye; when they are sketching, they use both eyes,
although it is not necessary to do so.
=Open the iris diaphragm=, and incline the mirror so that white light
is reflected on the Abbé condenser. Place the slide on the centre
of the stage, and if the slide contains a section of a plant, move
the slide so as to place this specimen over the centre of the Abbé
condenser. Then lower the body by means of the coarse adjustment
until the low-power object, which should always be in position when
work is begun, is within one-fourth of an inch of the stage. Then
raise the body by means of the coarse adjustment until the object,
or objects, in case a powder is being examined, is seen. Open and
close the iris diaphragm, finally adjusting the opening so that the
best possible illumination is obtained for bringing out clearly
the structure of the object or objects viewed. Then regulate the
focus by moving the body up or down by turning the fine adjustment.
When studying cross-sections or large particles of powders, it is
sometimes desirable to make low-power sketches of the specimen. In
most cases, however, only sufficient time should be spent in studying
the specimen to give an idea of the size, structure, and general
arrangement or plan or structure if a section of a plant, or, if
a powder, to note its striking characters. All the finer details
of structure are best brought out with the high-power objective in
position.
In =placing the high-power objective in position=, it is first
necessary to raise the body by the coarse adjustment; then open the
iris diaphragm, and lower the body until the objective is within
about one-eighth of an inch of the slide. Now raise the tube by
the fine adjustment until the object is in focus, then gradually
close the iris diaphragm until a clear definition of the object is
obtained. Now proceed to make an accurate sketch of the object or
objects being studied.
In =using the water or oil-immersion objectives= it is first
necessary to place a drop of distilled water or oil, as the case may
be, immediately over the specimen, then lower the body by the coarse
adjustment until the lens of the objective touches the water or the
oil. Raise the tube, regulate the light by the iris diaphragm, and
proceed as if the high-power objectives were in position.
The water or oil should be removed from the objectives and from the
slide when not in use.
After the higher-powered objective has been used, the body should
be raised, and the low-power objective placed in position. If the
draw-tube has been drawn out during the examination of the object,
replace it, but be sure to hold one hand on the nose-piece so as to
prevent scratching the objective and Abbé condenser by their coming
in forceful contact. Lastly, clean the mirror with a soft piece of
linen. In returning the microscope to its case, or to the shelf,
grasp the limb, or the pillar, firmly and carry as nearly vertical as
possible in order not to dislodge the eye-piece.
ILLUMINATION
The illumination for microscopic work may be from natural or
artificial sources.
[Illustration: FIG. 30.--Micro Lamp]
It has been generally supposed that the best possible illumination
for microscopic work is diffused sunlight obtained from a northern
direction. No matter from what direction diffused sunlight is
obtained, it will be found suitable for microscopic work. In no case
should direct sunlight be used, because it will be found blinding
in its effects upon the eyes. Natural illumination--diffused
sunlight--varies so greatly during the different months of the
year, and even during different periods of the day, that individual
workers are resorting more and more to artificial illumination. The
particular advantage of such illumination is due to the fact that
its quality and intensity are uniform at all times. There are many
ways of securing such artificial illumination, no one of which has
any particular advantage over the other. Some workers use an ordinary
gas or electric light with a color screen placed in the sub-stage
below the iris diaphragm. In other cases a globe filled with a weak
solution of copper sulphate is placed in such a way between the
source of light and the microscope that the light is focused on the
mirror. Modern mechanical ingenuity has devised, however, a number of
more convenient micro lamps (Fig. 30). These lamps are a combination
of light and screen. In some forms a number of different screens come
with each lamp, so that it is possible to obtain white-, blue-, or
dark-ground illumination. The type of the screen used will be varied
according to the nature of the object studied.
CARE OF THE MICROSCOPE
If possible, the microscope should be stored in a room of the same
temperature as that in which it is to be used. In any case, avoid
storing in a room that is cooler than the place of use, because when
it is brought into a warmer room, moisture will condense on the
ocular objectives and mirrors.
Before beginning work remove all moisture, dust, etc., from the inner
and outer lenses of the ocular, the objectives, the Abbé condenser,
and the mirror by means of a piece of soft, old linen. When the work
is finished the optical parts should be thoroughly cleaned.
If reagents have been used, be sure that none has got on the
objectives or the Abbé condenser. If any reagent has got on these
parts, wash it off with water, and then dry them thoroughly with soft
linen.
The inner lenses of the eye-pieces and the under lens of the Abbé
condenser should occasionally be cleaned. The mechanical parts of the
stand should be cleaned if dust accumulates, and the movable surfaces
should be oiled occasionally. Never attempt to make new combinations
of the ocular or objective lenses, or transfer the objectives or
ocular from one microscope to another, because the lenses of any
given microscope form a perfect lens system, and this would not be
the case if they were transferred. Keep clean cloths in a dust-proof
box. Under no circumstances touch any of the optical parts with your
fingers.
PREPARATION OF SPECIMENS FOR CUTTING
Most drug plants are supplied to pharmacists in a dried condition.
It is necessary, therefore, to boil the drug in water, the time
varying from a few minutes, in the case of thin leaves and herbs, up
to a half hour if the drug is a thick root or woody stem. If a green
(undried) drug is under examination, this first step is not necessary.
If the specimen to be cut is a leaf, a flower-petal, or other thin,
flexible part of a plant, it may be placed between pieces of elder
pith or slices of carrot or potato before cutting.
SHORT PARAFFIN PROCESS
In most cases, however, more perfect sections will be obtained if the
specimens are embedded in paraffin, by the quick paraffin process,
which is easily carried out.
After boiling the specimen in water, remove the excess of moisture
from the outer surface with filter paper or wait until the water has
evaporated. Next make a mould of stiff cardboard and pour melted
paraffin (melting at 50 or 60 degrees) into the mould to a height of
about one-half inch, when the paraffin has solidified. This may be
hastened by floating it on cool or iced water instead of allowing it
to cool at room temperature.
The specimens to be cut are now placed on the paraffin, with glue, if
necessary, to hold them in position, and melted paraffin poured over
the specimens until they are covered to a depth of about one-fourth
of an inch. Cool on iced water, trim off the outer paraffin to the
desired depth, and the Specimen will be in a condition suitable for
cutting.
Good workable sections may be cut from specimens embedded by this
quick paraffin method. After a little practice the entire process
can be carried out in less than an hour. This method of preparing
specimens for cutting will meet every need of the pharmacognosist.
LONG PARAFFIN PROCESS
In order to bring out the structure of the =protoplast= (living part
of the cell), it will be necessary to begin with the living part of
the plant and to use the long paraffin method or the collodion method.
Small fragments of a leaf, stem, or root-tip are placed in
chromic-acid solution, acetic alcohol, picric acid, chromacetic
acid, alcohol, etc., depending upon the nature of the specimen under
observation. The object of placing the living specimen in such
solutions is to kill the protoplast suddenly so that the parts of the
cell will bear the same relationship to each other that they did in
the living plant, and to fix the parts so killed.
After the fixing process is complete, the specimen is freed of the
fixing agent by washing in water. From the water-bath the specimens
are transferred successively to 10, 20, 40, 60, 70, 80, 90, and
finally 100 per cent alcohol. In this 100 per cent alcohol-bath the
last traces of moisture are removed. The length of time required to
leave the specimens in the different percentages of alcohols varies
from a few minutes to twenty-four hours, depending upon the size and
the nature of the specimen.
[Illustration: FIG. 31.--Paraffin-embedding Oven]
After dehydration the specimen is placed in a clearing
agent--chloroform or xylol--both of which are suitable when embedding
in paraffin. The clearing agents replace the alcohol in the cells,
and at the same time render the tissues transparent. From the
clearing agent the specimen is placed in a weak solution of paraffin,
dissolved xylol, or chloroform. The strength of the paraffin solution
is gradually increased until it consists of pure paraffin. The
temperature of the paraffin-embedding oven (Fig. 31) should not be
much higher than the melting-point of the paraffin.
The specimen is now ready to be embedded. First make a mould of
cardboard or a lead-embedding frame (Fig. 32), melt the paraffin, and
then place the specimen in a manner that will facilitate cutting.
Remove the excess of paraffin and cut when desired.
[Illustration: FIG. 32.--Paraffin Blocks]
In using the collodion method for embedding fibrous specimens,
as wood, bark, roots, etc., the specimen is first fixed with
picric acid, washed with water, cleared in ether-alcohol, embedded
successively in two, five, and twelve per cent ether-alcohol
collodion solution, and finally embedded in a pure collodion bath.
CUTTING SECTIONS
Specimens prepared as described above may be cut with a hand
microtome or a machine microtome.
HAND MICROTOME
In cutting sections by a =hand microtome=, it is necessary to place
the specimen, embedded in paraffin or held between pieces of elder
pith, carrot, or potato, over the second joints of the fingers,
then press the first joints firmly upon the specimen with the thumb
pressed against it. If they are correctly held, the specimens will be
just above the level of the finger and the end of the thumb, and the
joint will be below the level of the finger.
[Illustration: FIG. 33.--Hand Microtome]
Hold the section cutter (Fig. 33) firmly in the hand with the flat
surface next to the specimen. While cutting the section, press your
arm firmly against your chest, and bend the wrist nearly at right
angles to the arm. Push the cutting edge of the microtome toward the
body and through the specimen in such a way as to secure as thin a
section as possible. Do not expect to obtain nice, thin sections
during the first or second trials, but continued practice will enable
one to become quite efficient in cutting sections in this manner.
When the examination of drugs is a daily occurrence, the above method
will be found highly satisfactory.
MACHINE MICROTOMES
When a number of sections are to be prepared from a given specimen,
it is desirable to cut the sections on a machine microtome,
particularly when the sections are to be prepared for the use of
students, in which case they should be as uniform as possible.
Great care should be exercised in cutting sections with a machine
microtome--first, in the selection of the type of the microtome; and
secondly, in the style of knife used in cutting.
For soft tissues embedded in paraffin or collodion, the =rotary
microtome= with vertical knife will give best results. The thickness
of the specimen is regulated by mechanical means, so that in cutting
the sections it is only necessary to turn a crank and remove the
specimens from the knife-edge, unless there is a ribbon-carrier
attachment. If the sections are being cut from a specimen embedded
by the quick paraffin method, it is best to drop the section in a
metal cup partly filled with warm water. This will cause the paraffin
to straighten out, and the specimen will uncoil. After sufficient
specimens have been cut, the cup should be placed in a boiling-water
bath until the paraffin surrounding the sections melts and floats on
the water. Before removing the specimen from the water-bath, it is
advisable to shake the glass vigorously in order to cause as many
specimens as possible to settle to the bottom of the cup. The cup
is then placed in iced water or set aside until the paraffin has
solidified. The cake-like mass is then removed from the cup, and the
sections adhering to its under surface are removed by lifting them
carefully off with the flat side of the knife and transferring them,
together with the sections at the bottom of the cup, to a wide-mouth
bottle, and covered with alcohol, glycerine, and water mixture; or if
it is desired to stain the specimens, they should be placed in a weak
alcoholic solution.
Specimens having a hard, woody texture should be cut on a =sliding
microtome= by means of a special wood knife, which is especially
tempered to cut woody substances. Woody roots, wood, or thick bark
may be cut readily on this microtome when they have been embedded by
the quick paraffin process. The knife in the sliding microtome is
placed in a horizontal position, slanting so that the knife-edge is
drawn gradually across the specimen. After cutting, the sections are
treated as described above.
The thickness of the sections is regulated by mechanical means.
After a section has been cut, the block containing the specimen is
raised by turning a thumb-screw. In this microtome the knife, as in
the rotary type, is fixed, and the block containing the specimen is
movable.
If the specimen has been infiltrated with, and embedded in, paraffin
or collodion, the treatment of the sections after cutting should be
different.
In the case of paraffin, the sections are fastened directly to the
slide, and the paraffin is dissolved by either chloroform or xylol.
The specimen is then placed in 100, 95, and 45 per cent alcohol,
and then washed in water. These sections are now stained with
water-stains, brought back through alcohol, cleared, and mounted in
Canada balsam.
If alcoholic stains are used, it will not be necessary to dehydrate
before staining, and the dehydration after staining will also be
eliminated.
Sections infiltrated with collodion are either stained directly
without removing the collodion or after removal.
FORMS OF MICROTOMES
The =hand cylinder microtome= (Fig. 34) consists of a cylindrical
body. The clamp for holding the specimen is near the top below the
cutting surface. At the lower end is attached a micrometer screw with
a divided milled head. When moved forward one division, the specimen
is raised 0.01 mm. This micrometer screw has an upward movement of
10 mm. The cutting surface consists of a cylindrical glass ring.
[Illustration: FIG. 34.--Hand Cylinder Microtome]
[Illustration: FIG. 35.--Hand Table Microtome]
The =hand table microtome= (Fig. 35) is provided with a clamp, by
which it may be attached to the edge of a table or desk. The cutting
surface consists of two separated but parallel glass benches. The
object is held by a clamp and is raised by a micrometer screw, which,
when moved through one division by turning the divided head, raises
the specimen 0.01 mm.
The =sliding microtome= has a track of 250 mm. The object is held
by a clamp and its height regulated by hand. The disk regulating
the micrometer screw is divided into one hundred parts. When this
is turned through one division, the object is raised 0.005 mm. or
5 microns, at the same time a clock-spring in contact with teeth
registers by a clicking sound. If the disk is turned through two
divisions, there will be two clicks, etc. In this way is regulated
the thickness of the sections cut. When the micrometer screw has been
turned through the one hundred divisions, it must be unscrewed, the
specimen raised, and the steps of the process repeated. The knife is
movable and is drawn across the specimen in making sections.
[Illustration: FIG. 36.--Base Sledge Microtome]
The =base sledge microtome= (Fig. 36) has a heavy iron base which
supports a sliding-way on which the object-carrier moves. The
object-carrier is mounted on a solid mass of metal, and is provided
with a clamp for holding the object. The object is raised by turning
a knob which, when turned once, raises the specimen one to twenty
microns, according to how the feeding mechanism is set.
Sections thicker than twenty microns may be obtained by turning the
knob two or more times. The knife is fixed and is supported by two
pillars, the base of which may be moved forward or backward in such a
manner that the knife can be arranged with an oblique or right-angled
cutting surface.
[Illustration: FIG. 37.--Minot Rotary Microtome]
The =Minot rotary microtome= (Fig. 37) has a fixed knife, held in
position by two pillars, and a movable object-carrier. The object is
firmly secured by a clamp, and it is raised by a micrometer screw.
The screw is attached to a wheel having five hundred teeth on its
periphery. A pawl is adjusted to the teeth in such a way that, when
moved by turning a wheel to which it is attached, specimens varying
from one to twenty-five microns in thickness may be cut, according
to the way the adjusting disk is set. When the mechanism has been
regulated and the object adjusted for cutting, it is only necessary
to turn a crank in cutting sections.
CARE OF MICROTOMES
When not in use, microtomes should be protected from dust, and all
parts liable to friction should be oiled.
Microtome knives should be honed as often as is necessary to insure
a proper cutting edge. After cutting objects, the knives should be
removed, cleaned, and oiled.
It should be kept clearly in mind that special knives are required
for cutting collodion, paraffin, and frozen and woody sections. The
cutting edges of the different knives vary considerably, as is shown
in the preceding cuts.
CHAPTER V
REAGENTS
Little attention is given in the present work to micro-chemical
reactions for the reason that their value has been much overrated in
the past. A few reagents will be found useful, however, and these few
are given, as well as their special use. They are as follows:
LIST OF REAGENTS
=Distilled Water= is used in the alcohol, glycerine, and water
mixture as a general mounting medium. It is used when warm as a test
for inulin and it is used in preparing various reagents.
=Glycerine= is used in preparing the alcohol, glycerine, and water
mixture, in testing for aleurone grains, and as a temporary mounting
medium.
=Alcohol= is used in preparing the alcohol, glycerine, and water
mixture, in testing for volatile oils.
=Acetic Acid=. Both dilute and strong solutions are used in testing
for aleurone grains, cystoliths, and crystals of calcium oxalate.
=Hydrochloric Acid= is used in connection with phloroglucin as a test
for lignin and as a test for calcium oxalate.
=Ferric Chloride Solution= is used as a test for tannin.
=Sulphuric Acid= is used as a test for calcium oxalate.
=Tincture Alkana= is used when freshly prepared by macerating the
granulated root with alcohol and filtering, as a test for resin.
=Sodium Hydroxide=. A five per cent solution is used as a test for
suberin and as a clearing agent.
=Copper Ammonia= is used as a test for cellulose.
=Ammonical Solution of Potash= is used as a test for fixed oils.
The solution is a mixture of equal parts of a saturated solution of
potassium hydroxide and stronger ammonia.
=Oil of Cloves= is used as a clearing fluid for sections preparatory
to mounting in Canada balsam.
=Canada Balsam= is used as a permanent mounting medium for dehydrated
specimens, and as a cement for ringing slides.
=Paraffin= is used for general embedding and infiltrating.
=Lugol’s Solution= is used as a test for starch and for aleurone
grains and proteid matters.
=Osmic Acid=. A two per cent solution is used as a test for fixed
oils.
=Alcohol, Glycerine, and Water Mixture= is used as a temporary
mounting medium and as a qualitative test for fixed oils.
=Chlorzinc Iodide= is used as a test for suberin, lignin, cellulose,
and starch.
=Analine Chloride= is used as a test for lignified cell walls of bast
fibres and of stone cells.
=Phloroglucin=. A one per cent alcoholic solution is used in
connection with hydrochloric acid as a test for lignin.
=Hæmatoxylin-Delifields= is used as a test for cellulose.
[Illustration: FIG. 38.--Reagent Set]
REAGENT SET
Each worker should be provided with a set of =reagent bottles=
(Fig. 38). Such a set may be selected according to the taste of the
individual, but experience has shown that a 30 c.c. bottle with a
ground-in pipette and a rubber bulb is preferable to other types. In
such forms the pipettes are readily cleaned, and the rubber bulbs can
be replaced when they become old and brittle. The entire set should
be protected from dust by keeping it in a case, the cover of which
should be closed when the set is not in use.
MEASURING CYLINDER
In order accurately to measure micro-chemical reagents, it is
necessary to have a standard 50 c.c. cylinder (Fig. 39) graduated to
c.c.’s. Such a cylinder should form a part of the reagent set.
[Illustration: FIG. 39.--Measuring Cylinder]
STAINING DISHES
[Illustration: FIG. 40.--Staining Dish]
There is a great variety of =staining dishes= (Fig. 40), but for
general histological work a glass staining dish with groves for
holding six or more slides and a glass cover is most desirable.
CHAPTER VI
HOW TO MOUNT SPECIMENS
The method of procedure in mounting specimens for study varies
according to the nature of the specimen, its preliminary treatment,
and the character of the mount to be made. As to duration, mounts are
either temporary or permanent.
TEMPORARY MOUNTS
In preparing a =temporary mount=, place the specimen in the centre
of a clean slide and add two or more drops of the temporary mounting
medium, which may be water, or a mixture of equal parts of alcohol,
glycerine, and water, or some micro-chemical reagent, as weak Lugol’s
solution, solution of chloral hydrate, etc. Cover this with a cover
glass and press down gently. Remove the excess of the mounting
medium with a piece of blotting paper. Now place the slide on the
stage and proceed to examine it. Such mounts can of course be used
only for short periods of study; and when the period of observation
is finished, the specimen should be removed and the slide washed,
or the slide washing may be deferred until a number of such slides
have accumulated. At any rate, when the mounting medium dries, the
specimen is no longer suitable for observation.
PERMANENT MOUNTS
Permanent mounts are prepared in much the same way as temporary, but
of course the mounting medium is different. The kind of permanent
mounting medium used depends upon the previous treatment of the
specimen. If the specimen has been preserved in alcohol or glycerine
and water, it is usually mounted in glycerine jelly. If the specimen
in question is a powder, it is placed in the centre of the slide and
a drop or two of glycerine, alcohol, and water mixture added, unless
the powder was already in suspension in such a mixture. Cut a small
cube of glycerine jelly and place it in the centre of the powder
mixture. Lift up the slide by means of pliers, or grasp the two
edges between the thumb and finger and hold over a small flame of an
alcohol lamp, or place on a steam-bath until the glycerine jelly has
melted. Next sterilize a dissecting needle, cool, and mix the powder
with the glycerine jelly, being careful not to lift the point of the
needle from the slide during the operation. If the mixing has been
carefully done, few or no air-bubbles will be present; but if they
are present, heat the needle, and while it is white hot touch the
bubbles with its point, and they will disappear. Now take a pair of
forceps and, after securing a clean cover glass near the edge, pass
them three times through the flame of the alcohol lamp. While holding
it in a slanting position, touch one side of the powder mixture and
slowly lower the cover glass until it comes in complete contact with
the mixture. Now press gently with the end of the needle-handle, and
set it aside to cool. When it is cool, place a neatly trimmed label
on one end of the slide, on which write the name of the specimen,
the number of the series of which it is to form a part, etc. Any
excess of glycerine jelly, which may have been pressed out from the
edges of the cover glass, should not be removed at once, but should
be allowed to remain on the slide for at least one month in order to
allow for shrinkage due to evaporation. At the end of a month remove
the glycerine jelly by first passing the blade of a knife, held in
a vertical position, the back of the knife being next to the slide,
around the edge of the cover glass. After turning the knife-blade so
that the flat side is in contact with slide, remove the jelly outside
of the cover glass. Any remaining fragments should be removed with
a piece of old linen or cotton cloth. Finally, ring the edge of the
cover glass with microscopical cement, of which there are many types
to be had. If the cleaning has been done thoroughly, there is no
better ringing cement than Canada balsam.
In mounting cross-sections, the method of procedure is similar to the
above, with the exception that the glycerine jelly is placed at the
side of the specimen and not in the centre. While melting the jelly,
incline the slide in order to allow the melted glycerine jelly to
flow gradually over the specimen, thus replacing the air contained in
the cells and intercellular spaces. Finish the mounting as directed
above, but under no conditions should you stir the glycerine jelly
with the section.
If specimens, after having been embedded in paraffin or collodion,
are cut, cleared, stained, and dehydrated, they are usually mounted
in Canada balsam. A small drop of this substance, which may be
obtained in collapsible tubes, is placed at one side of the specimen.
While inclining the slide, gently heat until the Canada balsam covers
the specimen. Secure a cover glass by the aid of pliers, pass it
through the flame three times, and lower it slowly while holding it
in an inclined position. Press gently on the cover glass with the
needle-handle, and keep in a horizontal position for twenty-four
hours, then place directly in a slide box or cabinet, since no
sealing is required.
Glycerine is sometimes used to make permanent mounts, but it is
unsatisfactory, because the cover glass is easily removed and the
specimen spoiled or lost, unless ringed--a procedure which is not
easily accomplished. If the specimen is to be mounted in glycerine,
it must first be placed in a mixture of alcohol, glycerine, and
water, and then transferred to glycerine. Lactic acid is another
permanent liquid-mounting medium, which is unsatisfactory in the same
way as glycerine, but like glycerine, there are certain special cases
where it is desirable to use it. When this is used, the slides should
be kept in a horizontal position, unless ringed.
COVER GLASSES
Great care should be used In the selection of =cover glasses=,
however, not only as regards their shape but as to their thickness.
The standard tube length of the different manufacturers makes an
allowance of a definite thickness for cover glasses. It is necessary,
therefore, to use cover glasses made by the manufacturer of the
microscope in use.
Cover glasses are either square or round. Of each there are four
different thicknesses and two different sizes. The standard
thicknesses are: The small size is designated three-fourths and the
large size seven-eighths.
[Illustration: FIG. 41.--Round Cover Glass]
[Illustration: FIG. 42.--Square Cover Glass]
[Illustration: FIG. 43.--Rectangular Cover Glass]
=Cover glasses= are circular (Fig. 41), square (Fig. 42), or
rectangular (Fig. 43) pieces of transparent glass used in
covering the specimens mounted on glass slides. A few years ago
much difficulty was experienced in obtaining uniformly thick and
transparent cover glasses, but no such difficulty is experienced
to-day. The type of cover glass used depends largely upon the
character of the specimen to be mounted. The square and rectangular
glasses are selected when a series of specimens are to be mounted,
but in mounting powdered drugs and histological specimens the round
cover glasses are preferable because they are more sightly and more
readily cleaned and rinsed.
GLASS SLIDES
[Illustration: FIG. 44.--Glass Slide]
=Glass slides= (Fig. 44) are rectangular pieces of transparent glass
used as a mounting surface for microscopic objects. The slides are
usually three inches long by one inch wide, and they should be
composed of white glass, and they should have ground and beveled
edges. Slides should be of uniform thickness, and they should not
become cloudy upon standing.
SLIDE AND COVER-GLASS FORCEPS
Slides and cover glasses should be grasped by their edges. To the
beginner this is not easy. In order to facilitate holding slides and
cover glasses during the mounting process, one may use a slide and a
cover-glass =forceps=. The slide forceps consists of wire bent and
twisted in such a way that it holds a slide firmly when attached to
its two edges.
[Illustration: FIG. 45.--Histological Forceps]
[Illustration: FIG. 46.--Forceps]
[Illustration: FIG. 47.--Sliding-pin Forceps]
There are various forms of cover-glass holders, but only two types as
far as the method of securing the cover glass is concerned. First,
there are the bacteriological and the histological forceps (Fig. 45),
which are self-closing. The two blades of such forceps must be forced
apart by pressure in securing the cover glass. The second type of
forceps is that in which the two blades are normally separated (Fig.
46), it being necessary to press the blades to either side of the
cover glass in order to secure and hold it. There is a modification
of this type of forceps which enables one to lock the blades by means
of a sliding pin (Fig. 47), after the cover glass has been secured.
It is well to accustom oneself to one type, for by so doing one may
become dexterous in its use.
NEEDLES
[Illustration: FIG. 48.--Dissecting Needle]
Two =dissecting needles= (Fig. 48) should form a part of the
histologist’s mounting set. The handles may be of any material, but
the needle should be of tempered steel and about two inches long.
SCISSORS
[Illustration: FIG. 49.--Scissors]
Almost any sort of =scissors= (Fig. 49) will do for histology work,
but a small scissors with fine pointed blades, are preferred.
Scissors are useful in trimming labels and in cutting strips of
leaves and sections of fibrous roots that are to be embedded and cut.
SCALPELS
[Illustration: FIG. 50.--Scalpels]
=Scalpels= (Fig. 50) have steel blades and ebony handles. These vary
in regard to size and quality of material. The cheaper grades are
quite as satisfactory, however, as the more expensive ones, and for
general use a medium-sized blade and handle will be found most useful.
TURNTABLE
[Illustration: FIG. 51.--Turntable]
Much time and energy may be saved by ringing slides on a =turntable=
(Fig. 51). There is a flat surface upon which to rest the hand
holding the brush with cement, and a revolving table upon which the
slide to be ringed is held by means of two clips. In ringing slides,
it is only necessary to revolve the table, and at the same time to
transfer the cement to the edge of the cover glass from the brush
held in the hand.
LABELING
There are many ways of =labeling slides=, but the best method is to
place on the label the name of the specimen, the powder number, and
the box, the tray or cabinet number. For example:
Powdered Arnica Flowers
No. 80--Box A--600.
PRESERVATION OF MOUNTED SPECIMENS
[Illustration: FIG. 52.--Slide Box]
[Illustration: FIG. 53.--Slide Tray]
Accurately mounted, labeled, and ringed slides should be filed away
for future study and reference. Such =filing= may be done in slide
boxes, in slide trays, or in cabinets. Slide boxes are to be had
of a holding capacity varying from one to one hundred slides. For
general use, slide boxes (Fig. 52) holding one hundred slides will
be found most useful. Some workers prefer trays (Fig. 53), because
of the saving of time in selecting specimens. Trays hold twenty
slides arranged in two rows. The cover of the tray is divided into
two sections so that, if desired, only one row of slides is uncovered
at a time. Slide cabinets (Fig. 54) are particularly desirable for
storing large individual collections, particularly when the slides
are used frequently for reference. Large selections of slides should
be numbered and card indexed in order to facilitate finding.
[Illustration: FIG. 54.--Slide Cabinet]
Part II
TISSUES CELLS, AND CELL CONTENTS
CHAPTER I
THE CELL
The =cell= is the unit of structure of all plants. In fact the cell
is the plant in many of the lower forms--so called unicellular
plants. All plants, then, consist of one or more cells.
While cells vary greatly in size, form, color, contents, and
function, still in certain respects their structure is identical.
TYPICAL CELL
The typical vegetable cell is composed of a living portion or
=protoplast= and an external covering, or =wall=. The protoplast
includes everything within the wall. It is made up of a number of
parts, each part performing certain functions yet harmonizing with
the work of the cell as a whole. The protoplast (protoplasm) is a
viscid substance resembling the white of an egg. The protoplast, when
unstained and unmagnified, appears structureless, but when stained
with dyes and magnified, it is found to be highly organized. The
two most striking parts of the protoplast are the =cytoplasm= and
the =nucleus=. The part of the protoplast lining the innermost part
of the wall is the =ectoplast=, which is less granular and slightly
denser than most of the =cytoplasm=. The cytoplasm is decidedly
granular in structure.
In the cytoplasm occurs one or more cavities, =vacuoles=, filled with
=cell sap=. Embedded in the cytoplasm are numerous =chromatophores=,
which vary in color in the different cells, from colorless to yellow,
to red, and to green. The =nucleus= is the seat of the vital activity
of the cell, and the seat of heredity. The whole life and activity of
the cell centre, therefore, in and about the nucleus.
The outer portion of the nucleus consists of a thin
membrane or wall. The membrane encloses numerous granular
particles--=chromatin=--which are highly susceptible to organic
stains. Among the granules are thread-like particles or =linin=. Near
the centre of the nucleus are one or more small rounded nucleoli. The
liquid portion of the nucleus, filling the membranes and surrounding
the chromatin, linin, and nucleoli, is the =nuclear sap=.
Other cell contents characteristic of certain cells are crystals,
starch, aleurone, oil, and alkaloids. The detailed discussion of
these substances will be deferred until a later chapter.
The =cell wall= which surrounds the protoplast is a product of its
activity. The structure and composition of the wall of any given cell
vary according to the ultimate function of the cell. The walls may be
thin or thick, porous or non-porous, and colored or colorless. The
composition of cell walls varies greatly. The majority of cell walls
are composed of cellulose, in other cells of linin, in others of
cutin, and in still others of suberin, etc. In the majority of cells
the walls are laid down in a series of layers one over the other by
apposition, similar to the manner of building a pile of paper from
separate sheets. The first layer is deposited over the primary wall,
formed during cell division; to this is added another layer, etc. A
modification of this manner of growth is that in which the layers are
built up one over the other, but the building is gradually done by
the deposit of minute particles of cell-wall substance over the older
deposits. Such walls are never striated, as is likely to be the case
in cell walls formed by the first method. In other cells the walls
are increased in thickness by the deposition of new wall material in
the older membrane. The cell walls will be discussed more fully when
the different tissues are studied in detail.
INDIRECT CELL DIVISION (KARYOKINESIS)
The purpose of cell division is to increase the number of cells
of a tissue, an organ, an organism, or to increase the number of
organisms, etc. Such cell divisions involve, first, an equal division
of the protoplast and, secondly, the formation of a wall between
the divided protoplasts. The first changes in structure of a cell
undergoing division occur in the nucleus.
CHANGES IN A CELL UNDERGOING DIVISION
The =linin threads= become thicker and shorter. The =chromatin
granules= increase in size and amount; the threads and chromatin
granules separate into a definite number of segments or =chromosomes=
(Plate 1, Fig. 2). The nuclear membrane becomes invested with a
fibrous protoplasmic layer which later separates and passes into
either end of the cell, there forming the =polar caps= (Plate 1, Fig.
3).
The =nuclear membrane= and the =nucleoli= disappear at about this
time. Two fibres, one from each polar cap, become attached to
opposite sides of the individual chromosomes. Other fibres from
the two polar caps unite to form the =spindle fibres=, which thus
extend from pole to pole. All these spindle fibres form the =nuclear
spindle= (Plate 1, Fig. 5).
The chromosomes now pass toward the division centre of the cell or
=equatorial plane= and form, collectively, the =equatorial plate=
(Plate 1, Fig. 5). At this point of cell division, the chromosomes
are =U=-shaped, and the curved part of the chromosomes faces the
equatorial plane. The chromosomes finally split into two equal
parts (Plate 1, Fig. 6). The actual separation of the halves of
chromosomes is brought about by the attached polar fibres, which
contract toward the polar caps (Plate 1, Fig. 7). The chromosomes are
finally drawn to the polar caps (Plate 1, Fig. 8). The chromosomes
now form a rounded mass. They then separate into linin threads
and chromatin granules. Nucleoli reappear, and nuclear sap forms.
Finally, a nuclear membrane develops. The spindle fibres, which still
extend from pole to pole, become thickened at the equatorial plane
(Plate 1, Fig. 8), and finally their edges become united to form the
=cell-plate= (Plate 1, Fig. 9), which extends across the cell, thus
completely separating the mother cell into two daughter cells. After
the formation of the cell-plate, the spindle fibres disappear. The
cell becomes modified to form the =middle lamella=, on either side of
which the daughter protoplast adds a cellulose layer. The ultimate
composition of the middle lamella and the composition and structure
of the cell wall will differ according to the function which the cell
will finally perform.
[Illustration: PLATE 1
Nine figures, showing stages in the cell-division common to the
onion root (_Allium cepa_, L.)]
ORIGIN OF MULTICELLULAR PLANTS
All multicellular plants are built up by the repeated cell division
of one original cell. If the cells formed are similar in structure
and function, they form a tissue. In multicellular plants many
different kinds of tissues will be formed as a result of cell
division, since there are many different functions to be performed by
such an organism. When several of these tissues become associated and
their functions are correlated, they form an organ. The association
of several organs in one form makes an organism. The oak-tree is an
organism. It is made up of organs known as flowers, leaves, stems,
roots, etc. Each of these organs is in turn made up of several kinds
of tissue. In some cases it is difficult to designate a single
function to an aggregation of cells (tissue). In fact, a tissue may
perform different functions at different periods of its existence
or it may perform two functions at one and the same time; as an
example, stone cells, whose primary function is mechanical, in many
cases function as storage tissue. The cells forming the tissues of
the plant, in fact, show great adaptability in regard to the function
which they perform. Nevertheless there is a predominating function
which all tissues perform, and the structure of the cells forming
such tissues is so uniform that it is possible to classify them.
The functional classification of tissues is chosen for the purpose
of demonstrating the adaptation of cell structure to cell function.
If the cells performing a similar function in the different plants
were identical in number, distribution, form, color, size, structure,
and cell contents, there would not be a science of histology upon
which the art of microscopic pharmacognosy is based. It may be said,
however, with certainty, that the cells forming certain of the
tissues of any given species of plant will differ in a recognizable
degree from cells performing a similar function in other species of
plants. Often a tissue is present in one plant but absent in another.
For example, many aquatic plants are devoid of mechanical fibrous
cells. The barks of certain plants have characteristic stone cells,
while in many other barks no stone cells occur. Many leaves have
characteristic trichomes; others are free from trichomes, etc. Yet
all cells performing a given function will structurally resemble
each other. In the present work the nucleus and other parts of the
living protoplast will not be considered, for the reason that these
parts are not in a condition suitable for study, because most drugs
come to market in a dried condition, a condition which eliminates
the possibility of studying the protoplast. The general structure of
the cells forming the different tissues will first be considered,
then their variation, as seen in different plants, and finally their
functions.
CHAPTER II
THE EPIDERMIS AND PERIDERM
The epidermis and its modifications, the hypodermis and the periderm,
form the dermal or protective outer layer or layers of the plant.
The epidermis of most leaves, stems of herbs, seeds, fruits, floral
organs, and young woody stems consists of a single layer of cells
which form an impervious outer covering, with the exception of the
stoma.
LEAF EPIDERMIS
The cells of the =epidermis= vary in size, in thickness of the side
and end walls, in form, in arrangement, in character of outgrowths,
in the nature of the surface deposits, in the character of
wall--whether smooth or rough--and in size.
In cross-sections of the leaf the character of both the side and end
walls is easily studied.
In surface sections--the view most frequently seen in powders--the
side walls are more conspicuous than the end wall (Plates 2 and 3).
This is so because the light is considerably retarded in passing
through the entire length of the side walls, while the light is
retarded only slightly in passing through the end wall. The light in
this case passes through the width (thickness) of the wall only. The
outer walls of epidermal cells are characteristic only when they are
striated, rough, pitted, colored, etc. In the majority of leaves the
outer wall of the epidermal cells is not diagnostic in powders, or in
surface sections.
The thickness of the end and side walls of epidermal cells differs
greatly in different plants.
As a rule, leaves of aquatic and shade-loving plants, as well as the
leaves of most herbs have thinner walled epidermal cells than have
the leaves of plants growing in soil under normal conditions, or than
have the leaves of shrubs and trees.
[Illustration: PLATE 2
LEAF EPIDERMIS
1. Uva-ursi (_Arctostaphylos uva-ursi_, [L.] Spring).
2. Boldus (_Peumus boldus_, Molina).
3. Catnip (_Nepeta cataria_, L.).
4. Digitalis (_Digitalis purpurea_, L.).
4-A. Origin of hair.]
[Illustration: PLATE 3
LEAF EPIDERMIS
1. Upper striated epidermis of chirata leaf (_Swertia chirata_,
[Roxb.] Ham.).
2. Green hellebore leaf (_Veratrum viride_, Ait.).
3. Boldus leaf (_Peumus boldus_, Molina).
4. Under epidermis of India senna (_Cassia angustifolia_, Vahl.).]
The widest possible range of cell-wall thickness is therefore found
in the medicinal leaves, because the medicinal leaves are collected
from aquatic plants, herbs, shrubs, trees, etc.
The outer wall is always thicker than the side walls. Even the
side walls vary in thickness in some leaves, the wall next to the
epidermis being thicker than the lower or innermost portion of the
wall. Frequently the outermost part of the side walls is unequally
thickened. This is the case in the beaded side walls characteristic
of the epidermis of the leaves of laurus, myrcia, boldus, and
capsicum seed, etc. The thickness of the side walls of the epidermal
cells of most leaves varies in the different leaves.
In most leaves there are five typical forms of arrangement of
epidermal cells: First, those over the veins which are elongated
in the direction of the length of the leaf; and, secondly, those
on other parts of the leaf which are usually several-sided and not
elongated in any one direction. If the epidermis of the leaf has
stoma, then there is a third type of arrangement of the epidermal
cells around the stoma; fourthly, the cells surrounding the base of
hairs; and fifthly, outgrowths of the epidermis, non-glandular and
glandular hairs, etc.
It should be borne in mind that in each species of plant the five
types of arrangement are characteristic for the species.
The character of the outer wall of the epidermal cells differs
greatly in different plants. In most cases the wall is smooth; senna
is an example of such leaves. In certain other leaves the wall
is rough, the roughness being in the form of striations. In some
cases the striations occur in a regular manner; belladonna leaf is
typical of such leaves. In other instances the wall is striated in
an irregular manner as shown in chirata epidermis. Very often an
epidermis is rough, but the roughness is not due to striations.
In these cases the epidermis is unevenly thickened, the thin
places appearing as slight depressions, the thick places as slight
elevations. Boldus has a rough, but not a striated surface.
=Surface deposits= are not of common occurrence in medicinal plants;
waxy deposits occur on the stem of sumac, on a species of raspberry,
on the fruit of bayberry, etc. Resinous deposits occur on the leaves
and stems of grindelia species, and on yerba santa.
In certain leaves there are two or three layers of cells beneath the
epidermis that are similar in structure to the epidermal cells. These
are called hypodermal cells, and they function in the same way as the
epidermal cells.
Hypodermal cells are very likely to occur on the margin of the leaf.
Uva-ursi leaf has a structure typical of leaves with hypodermal
marginal cells. Uva-ursi, like other leaves with hypodermal cells has
a greater number of hypodermal cells at the leaf margin than at any
other part of the leaf surface.
The cutinized walls of epidermal cells are stained red with saffranin.
TESTA EPIDERMIS
=Testa epidermal cells= form the epidermal layers of such seeds as
lobelia, henbane, capsicum, paprika, larkspur, belladonna, scopola,
etc.
In surface view the end walls are thick and wavy in outline;
frequently the line of union--middle lamella--of two cells is
indicated by a dark or light line, while in others the wall between
two cells appears as a single wall. The walls are porous or
non-porous, and the color of the wall varies from yellow to brown, to
colorless. These cells always occur in masses, composed partially of
entire and partially of broken fragments.
In lobelia seed (Plate 4, Fig. 2) the line of union of adjacent cell
walls appears as a dark line. The walls are wavy in outline, of a
yellowish-red color and not porous.
In henbane seed (Plate 4, Fig. 3) the line of union between the cells
is scarcely visible; the walls are decidedly wavy, more so than in
lobelia, and no pits are visible.
In capsicum seed (Plate 4, Fig. 1) the cells are very wavy and
decidedly porous, the line of union between the cell walls being
marked with irregular spaces and lines.
In belladonna seed (Plate 5, Fig. 1) the walls between two adjacent
cells are non-striated and non-porous, and extremely irregular in
outline.
[Illustration: PLATE 4
TESTA EPIDERMAL CELLS
1. Capsicum seed (_Capsicum frutescens_, L.).
2. Lobelia seed (_Lobelia inflata_, L.).
3. Henbane seed (_Hyoscyamus niger_, L.).]
[Illustration: PLATE 5
TESTA CELLS
1. Belladonna seed (_Atropa belladonna_, L.).
2. Star-aniseed (_Illicium verum_, Hooker).
3. Stramonium seed (_Datura stramonium_, L.).]
In star-anise seed (Plate 5, Fig. 2) the walls are irregularly
thickened and wavy in outline.
In stramonium seed (Plate 5, Fig. 3) the walls are very thick, wavy
in outline, and striated.
PLANT HAIRS (TRICHOMES)
In histological work plant hairs are of great importance, as they
offer a ready means of distinguishing and differentiating between
plants, or parts of plants, when they occur in a broken or finely
powdered condition. There is no other element in powdered drugs which
is of so great a diagnostic value as the plant hair. The same plant
will always have the same type of hair, the only noticeable variation
being in the size. In microscopical drug analysis the presence of
hairs is always noted, and in many cases the purity of the powder
can be ascertained from the hairs. Botanists seem to have given
little attention to the study of plant hairs. This accounts for the
fact that information concerning them is very meagre in botanical
literature, and, as far as the author can learn, no one has attempted
to classify them. In systematic work, plant hairs could be used to
great advantage in separating genera and even species. Hairs are,
of course, a factor now in systematic work. The lack of hairs is
indicated by the term glabrous. Their presence is indicated by such
terms as hispid, villous, etc. In certain cases the term indicates
position of the hair as ciliate when the hair is marginal. When hairs
influence the color of the leaf, such terms as cinerous and canescent
are used. In all the cases cited no mention is made of the real
nature of the hair.
In systematic work, as in pharmacognosy, we must work with dried
material, and it is only those hairs which retain their form under
such conditions which are of classification value.
Hairs are the most common outgrowths of the epidermal cells. They
are classified as glandular or non-glandular, according to their
structure and function. The glandular hairs will be considered under
synthetic tissue.
Each group is again subdivided into a number of secondary groups,
depending upon the number of cells present, their form, their
arrangement, their size, their color, the character of their walls,
whether rough or smooth, whether branched or non-branched, whether
curved, twisted, straight, or twisted and straight, whether pointed,
blunt, or forked.
FORMS OF HAIRS
PAPILLÆ
=Papillæ= are epidermal cells which are extended outward in the form
of small tubular outgrowths.
Papillæ occur on the following parts of the plant: flower-petals,
stigmas, styles, leaves, stems, seeds, and fruits. Papillæ occur on
only a few of the medicinal leaves.
The under surface of both Truxillo (Plate 6, Fig. 3) and Huanuca coca
have very small papillæ. The outermost wall of these papillæ are much
thicker than the side walls. The papillæ of klip buchu (Plate 6, Fig.
4), an adulterant of true buchu, has large thick-walled papillæ.
The velvety appearance of most flower-petals (Plate 6, Figs. 2 and
5) is due to the presence of papillæ. The papillæ of flower-petals
are very variable. In calendula flowers (Plate 6, Fig. 1) they
are small, yellowish in color, and the outer wall is marked with
parallel striations which appear as small teeth in cross-section.
The ray petal papillæ of anthemis consist of rather large, broad,
blunt papillæ with slightly striated walls. The papillæ of the ray
petals of the white daisy consist of papillæ which have medium sized,
cone-shaped papillæ with finely striated walls. The papillæ of the
flower stigma vary greatly in different flowers. In some cases two or
more types of papillæ occur, but even in these cases the papillæ are
characteristic of the species.
The papillæ differ greatly in the case of the flowers of the
compositæ, where two types of flowers are normally present--namely,
the ray flowers and the disk flowers.
In all cases observed the papillæ of the stigma of the ray flowers
are always smaller than the papillæ of the stigma of the disk
flowers. It would appear from extended observation that the papillæ
of the ray flower stigma are being gradually aborted. The papillæ of
the style are always different from the papillæ of the stigma. The
style papillæ are always smaller, and they are of a different form.
[Illustration: PLATE 6
PAPILLÆ
1. Calendula flowers (_Calendula officinalis_, L.).
2. White daisy ray flower (_Chrysanthemum leucanthemum_, L.).
3. Coca leaf (_Erythroxylon coca_, Lamarck).
4. Klip buchu.
5. Anthemis ray petal (_Anthemis nobilis_, L.).]
UNICELLULAR NON-GLANDULAR HAIRS
=True plant hairs= are tubular outgrowths of the epidermal cell, the
length of these outgrowths being several times the width of the hair.
The unicellular hairs are common to many plants. The two groups
of non-glandular unicellular hairs are, first, the solitary; and
secondly, the clustered hairs.
=Solitary unicellular hairs= occur on the leaves of chestnut, yerba
santa, lobelia, cannabis indica, the fruit of anise, and the stem of
allspice, senna, and cowage.
Chestnut hairs (Plate 7, Fig. 1) have smooth yellowish-colored walls,
and the cell cavity contains reddish-brown tannin. These hairs occur
solitary or clustered; the clustered hairs normally occur on the
leaf, but in powdering the drug, individual hairs of the cluster
become separated or solitary.
Yerba santa hairs (Plate 7, Fig. 4) are twisted, the lumen or cell
cavity is very small, and the walls, which are very thick, are
grayish-white.
Lobelia hairs (Plate 7, Fig. 5) are very large. The walls are
grayish-white, and the outer surface extends in the form of small
elevations which make the hair very rough. The hair tapers gradually
to a solid point.
Cannabis indica hairs (Plate 7, Fig. 6) are curved. The apex tapers
to a point and the base is broad, and it frequently contains deposits
of calcium carbonate. The walls are grayish-white in appearance, and
rough. The roughness increases toward the apex.
The hairs of anise (Plate 7, Fig. 7) are mostly curved; the walls are
thick, yellowish-white, and the outer surface is rough; this is due
to the numerous slight centrifugal projections of the outer wall.
Allspice stem hairs (Plate 7, Fig. 2) have smooth walls. The cell
cavity is reddish-brown. The hair is curved.
The hair of senna (Plate 7, Fig. 10) is light greenish-yellow with
rough papillose walls. The hair is usually curved and tapering, and
it does not have any characteristic cell contents.
[Illustration: PLATE 7
UNICELLULAR SOLITARY HAIRS
1. Chestnut leaf (_Castanea dentata_, [Marsh] Borkh).
2. Allspice stems (_Pimento, officinalis_, Lindl.).
3. Cowage.
4. Yerba santa (_Eriodictyon californicum_, [H. and A.] Greene).
5. Lobelia (_Lobelia inflata_, L.).
6. Cannabis indica (_Cannabis saliva_, L.).
7. Anise fruit (_Pimpinella anisum_, L.).
8. Hesperis matronalis (_Hesperis matronalis_, L.).
9. Galphimia glauca (_Galphimia glauca_, Cav.).
10. Senna (_Cassia angustifolia_, Vahl.).]
[Illustration: PLATE 8
CLUSTERED UNICELLULAR HAIRS
1. and 2. European oak (_Quercus infectoria_, Olivier).
3. Kamala (_Mallotus philippinensis_, [Lam.] [Muell.] Arg.).
4. Witch-hazel leaf (_Hamamelis virginiana_, L.).
5. Althea leaf (_Althæa officinalis_, L.).]
Cowage hairs (Plate 7, Fig. 3) are lance-shaped, and they
terminate in a sharp point. The outer wall contains numerous
recurved teeth-like projections. The cell cavity is filled with a
reddish-brown contents which are somewhat fissured.
=Clustered unicellular hairs= occur on the leaves of chestnut,
witch-hazel, althea, European oak, etc. In European oak (Plate 8,
Figs. 1 and 2) clusters of two and three hairs occur. The walls are
yellowish-white, smooth, and the tip of the hair is solid.
In kamala (Plate 8, Fig. 3) clusters of seven or more hairs occur;
the walls are yellowish, and the cell cavity is reddish-brown. In
witch-hazel leaf (Plate 8, Fig. 4) clusters of a variable number
of hairs occur. The hairs, which are of various lengths, have
yellowish-white, thick, smooth walls, and reddish cell contents.
In althea leaf (Plate 8, Fig. 5) the hairs are nearly straight and
the walls are smooth. The basal portions of the hair are strongly
pitted.
=Branched solitary unicellular hairs= occur on the leaves of hesperis
matronalis (Plate 7, Fig. 8), and on galphimia glauca (Plate 7, Fig.
9).
The hair of hesperis matronalis has smooth walls, and the two
branches grow out nearly parallel to the leaf surface.
The hair of galphimia glauca has rough walls, and the two branches
grow upward in a bifurcating manner.
MULTICELLULAR HAIRS
=Multicellular= hairs are divided into the uniseriate and the
multiseriate hairs. Both of these groups are divided into the
branched and the non-branched hairs, as follows:
1. =Uniseriate=.
(_A_) =Non-branched.=
(_B_) =Branched.=
2. =Multiseriate.=
(_A_) =Non-branched.=
(_B_) =Branched.=
=Multicellular uniseriate non-branched hairs= occur on the leaves of
digitalis, Western and Eastern skullcap, peppermint, thyme, yarrow,
arnica flowers, and sumac fruit.
[Illustration: PLATE 9
MULTICELLULAR UNISERIATE NON-BRANCHED HAIRS
1. Digitalis leaf (_Digitalis purpurea_, L.).
2. Arnica flower (_Arnica montana_, L.).
3. Western skullcap plant (_Scutellaria canescens_, Nutt.).
4. Eastern skullcap plant (_Scutellaria lateriflora_, L.).
5. Peppermint leaf (_Mentha piperita_, L.).
6. Thyme leaf (_Thymus vulgaris_, L.).
7. Yarrow flowers (_Achillea millefolium_, L.).
8. Wormwood leaf (_Artemisia absinthium_, L.).
9. Sumac fruit (_Rhus glabra_, L.).]
Digitalis hairs (Plate 9, Fig. 1) are made up of a varying number of
uniseriate-arranged cells of unequal length, frequently placed at
right angles to the cells above and below; the walls are of a whitish
color, and are rough or smooth.
Eastern skullcap (Plate 9, Fig. 4) has hairs with not more than four
cells; these hairs are curved, and the walls are whitish, sometimes
smooth, but usually rough. In Western skullcap (Plate 9, Fig. 3) the
hairs have sometimes as many as seven cells. The walls are white and
rough, and the individual cells of the hair are much larger than are
the cells of the hairs of true skullcap.
Peppermint (Plate 9, Fig. 5) has from one to eight cells. The hair is
curved, and the walls are very rough.
Thyme (Plate 9, Fig. 6) has short, thick, rough-walled trichomes, the
terminal cell usually being bent at nearly right angles to the other
cells.
Yarrow hairs (Plate 9, Fig. 7) have a variable number of cells. In
all the hairs the basal cells are short and broad, while the terminal
cell is greatly elongated.
Arnica hairs (one form, Plate 9, Fig. 2) have frequently as many as
four cells, the terminal cell being longer than the basal cells. The
walls are white and smooth.
Sumac-fruit hairs (Plate 9, Fig. 9) have spindle-shaped,
reddish-colored hairs.
=Multicellular multiseriate non-branched hairs= occur on cumin fruit
and on the tubular part of the corolla of calendula.
The hairs on cumin fruit vary considerably in size. All the hairs
are spreading at the base and blunt or rounded at the apex. The
cells forming the hair are narrow and the walls are thick. Three
differently sized hairs are shown in Plate 10, Fig. 1.
The hairs of the base of the ligulate petals of calendula (Plate 10,
Fig. 2) are biseriate. The hairs are very long and the walls are very
thin.
=Multicellular uniseriate branched hairs= occur on the leaves of
dittany of Crete, mullen, and on the calyx of lavender flowers.
The dittany of Crete (Plate 11, Fig. 3) hair is smooth-walled, and
the branches are alternate.
In mullen (Plate 11, Fig. 1) the hairs have whorled branches, the
walls are smooth, and the cell cavity usually contains air.
[Illustration: PLATE 10
MULTICELLULAR MULTISERIATE NON-BRANCHED HAIRS
1. Cumin (_Cuminum cyminum_, L.).
2. Marigold (_Calendula officinalis_, L.).]
[Illustration: PLATE 11
MULTICELLULAR UNISERIATE BRANCHED HAIRS
1. Mullen leaf (_Verbascum thapsus_, L.).
2. Lavender flowers (_Lavandula vera_, D. C.).
3. Dittany of Crete (_Origanum dictamnus_, L.).]
The lavender hairs (Plate 11, Fig. 2) have mostly opposite branches,
and the walls are rough. Thus the multicellular branched hairs may be
divided into subgroups which have alternate, opposite, whorled, or in
certain hairs irregularly arranged branches. Each class may be again
subdivided according to color, character of cell termination, etc.,
as cited at the beginning of the chapter.
Occasionally multicellular hairs assume the form of a shield (Plate
12, Fig. 1); in such cases the hair is termed peltate, as in the
non-glandular multicellular hair of shepherdia canadensis.
Hairs grow out from the surface of the epidermis in a perpendicular,
a parallel, or in an oblique direction. Hairs which grow parallel or
oblique to the surface are usually curved, and the outer curved part
of the wall is usually thicker than the inner curved wall.
The mature hairs of some plants consist of dead cells. In other
plants the cells forming the hair are living. When dried, those
hairs, which were dead before drying, contain air; while those hairs
which were living before drying, show great variation in color and in
the nature of the cell contents. The contents are either organic or
inorganic. The commonest organic constituent is dried protoplasm. In
cannabis indica are deposits of calcium carbonate.
=Multicellular multiseriate branched hairs= are the ultimate division
of the pappus of erigeron, aromatic goldenrod, arnica, grindelia,
boneset, and life-everlasting.
The hairs of erigeron (Plate 13, Figs. 1 and 2) are slender; the
walls are porous. Each hair terminates in two cells, which are
greatly extended and sharp-pointed; the branches from the basal part
of the hairs (Plate 13, Fig. 1) are of about the same length as the
apical branches.
The hairs of aromatic goldenrod (Plate 13, Figs. 3 and 4) are larger
than those of erigeron; the diameter is greater and the walls are
non-porous. The apex of the hair terminates in a group of about four
cells of unequal length, which are sharp-pointed. The branches of the
basal cells (Plate 13, Fig. 3) are similar to the branches of the
apical cells.
The hairs of arnica (Plate 14, Figs. 1 and 2) have thick, strongly
porous walls; the branches terminate in sharp points. The apex of the
hair terminates in a single cell. The basal branches (Plate 14, Fig.
2) are much longer than special branches.
[Illustration: PLATE 12
NON-GLANDULAR MULTICELLULAR HAIRS
_Shepherdia canadensis_, [L.] Nutt.]
[Illustration: PLATE 13
MULTICELLULAR MULTISERIATE BRANCHED HAIRS
1. Basal hairs of erigeron (_Erigeron canadensis_, L.).
2. Apical hairs of erigeron (_Erigeron canadensis_, L.).
3. Basal hairs of aromatic goldenrod (_Solidago odora_, Ait.).
4. Apical hairs of aromatic goldenrod (_Solidago odora_, Ait.).]
The hair of grindelia (Plate 14, Figs. 3 and 4) has very thick walls
with numerous elongated pores. The apex of the hair terminates in
a cluster of cells with short, free, sharp-pointed ends. The basal
branches (Plate 14, Fig. 4) are longer than the apical branches.
Boneset hair (Plate 15, Figs. 1 and 2) has non-porous walls. The
apex of the hair terminates in two blunt-pointed cells. The terminal
wall is thicker than the side wall. Some of the branches lower
down terminate in cells with very thick or solid points. The basal
branches (Plate 15, Fig. 1) are longer, but the cells are narrower
and more strongly tapering than are the branches of the apical part
of the hair.
Life-everlasting (Plate 15, Figs. 3 and 4) has uniformly thickened
but non-porous walls. The hair terminates in two blunt-pointed,
greatly elongated cells.
The basal branches (Plate 15, Fig. 4) are narrower, slightly
tapering, and the base of the branches frequently curve downward.
The cell cavities of these hairs are filled with air.
The walls of hairs are composed of cutin, of lignin, and of cellulose.
PERIDERM
The =periderm= is the outer protective covering of the stems
and roots of mature shrubs and trees. The periderm replaces the
epidermis. The periderm may be composed of cork cells, stone
cell-cork, or a mixture of cork, parenchyma, fibres, stone cells, etc.
CORK PERIDERM
The typical periderm is made up of =cork cells=. Cork cells vary in
appearance, according to the part of the cell viewed.
[Illustration: PLATE 14
MULTICELLULAR MULTISERIATE BRANCHED HAIRS
1. Apical hairs arnica (_Arnica montana_, L.).
2. Basal hairs arnica (_Arnica montana_, L.).
3. Apical hairs grindelia (_Grindelia squarrosa_, [Pursh] Dunal).
4. Basal hairs grindelia (_Grindelia squarrosa_, [Pursh] Dunal).]
[Illustration: PLATE 15
MULTICELLULAR MULTISERIATE BRANCHED HAIRS
1. Apical hairs boneset (_Eupatorium perfoliatum_, L.).
2. Basal hairs boneset (_Eupatorium perfoliatum_, L.).
3. Apical hairs life-everlasting (_Gnaphalium obtusifolium_, L.).
4. Basal hairs life-everlasting (_Gnaphalium obtusifolium_, L.).]
On surface view (Plate 16, Fig. A) the cork cells are angled in
outline and are made up of from four to seven side walls; five-
and six-sided cells are more common than the four-and seven-sided
cells. Surface sections of cork cells show their length and width.
These side walls usually appear nearly white, while the end wall,
particularly of the outermost cork cells, usually appears brown or
reddish-brown, or in some cases nearly black.
Cork cells on cross-section are rectangular in form, and they are
arranged in superimposed rows, the number of rows being gradually
increased as the plant grows older. Such an increase in the number of
rows of cork cells is shown in the cross-section of cascara sagrada
(Plate 16, Fig. C).
Cork cells fit together so closely that there is no intercellular
spaces between the cells. In this case two rows of cork cells occupy
no greater space than the solitary row of cork cells immediately over
and external to them. As a rule, the outermost layers of cork cells
have a narrower radial diameter than the cork cells of the underlying
layers. This is due to the fact that these outer cells are stretched
as the stem increases in diameter. This view shows the height of
cork cells, but not always the length, which will, of course, vary
according to the part of the cell cut across. In a section a few
millimeters in diameter, however, all the variations in size may be
observed. The color of the walls is nearly white.
The cavity may contain tannin or other substances. When tannin is
present, the cavity is of a brownish or brownish-red color, or it may
be nearly black. Most barks appear devoid of any colored or colorless
cell contents.
The radial section (Plate 16, Fig. B) of cork cells shows the height
of the cells and the width of the cells at the point cut across. Some
cells will be cut across their longest diameter, while others will
be cut across their shortest diameter. Cork cells are, therefore,
smaller in radial section than they are in cross-section. The color
of the walls is white, and the color and nature of the cell contents
vary for the same reasons that they vary in cross-sections.
The number of layers of cork cells occurring in cross- and
radial-sections varies according to the age of the plant, to the type
of plant, and to the conditions under which the plant is growing.
The number of layers of cork cells is not of diagnostic importance,
nor is the surface view of cork cells diagnostic except in certain
isolated cases.
[Illustration: PLATE 16
PERIDERM OF CASCARA SAGRADA (_Rhamnus purshiana_, D.C.)
_A._ 1, Outline of cork cells; 2, Line of contact of adjoining cork
cells.
_B._ Radial longitudinal section of cascara sagrada. 1, Cork cells;
2, Phellogen; 3, Forming parenchyma cells; 4, Cortical parenchyma
cells.
_C._ Cross-section of cascara sagrada. 1, Cork cells; 2, Phellogen;
3, Forming parenchyma cells; 4, Cortical parenchyma cells.]
The presence or absence of cork or epidermal tissue in powders must
always be noted. The presence of cork enables one to distinguish
Spanish from Russian licorice. In like manner, the presence of
epidermis enables one to distinguish the pharmacopœial from the
unofficial peeled calamus. The absence of epidermis in Jamaica ginger
is one of the means by which this variety is distinguished from the
other varieties of ginger, etc.
In canella alba the periderm is replaced by stone cell-cork. That is,
the cells forming the periderm are of a typical cork shape, but the
walls are lignified, unequally thickened, and the inner or thicker
walls are strongly porous, and the walls are of a yellowish color.
Stone cell-cork forms the periderm of clove bark also, but the cells
are narrower and longer, and the inner wall is not so thick or porous
as is the case in canella alba bark.
STONE CELL PERIDERM
In canella alba (Plate 17, Fig. B) cork periderm is frequently
replaced by stone cells, particularly in the older barks. These stone
cells form the periderm because they replace the cork periderm, which
fissures and scales off as the root increases in diameter.
The side and end walls of cork cells are of nearly uniform diameter.
Exceptions occur, but they are not common. In buchu stem (Plate
101, Fig. 3), the cork cells have thick outer walls, but thin sides
and inner walls. The cell cavity contains reddish-brown deposits of
tannin.
PARENCHYMA AND STONE CELL PERIDERM
As the trees and shrubs increase in diameter, cracks or fissures
occur in the periderm, or corky layer. In such cases the phellogen
cells divide and redivide in such manner as to cut off a portion of
the parenchyma cells, stone cells, and fibres of the cortex which
is inside of and below the fissure. All the parenchyma cells, etc.,
exterior to the newly formed cork cells soon lose their living-cell
contents, since their food-supply is cut off by the impervious walls
of the cork cells. In time they are forced outward by the developing
cork cells until they partially or completely fill the break in the
periderm. In white oak bark (Plate 18), as in other barks, a large
part of the periderm is composed of dead and discolored cortical
cells.
[Illustration: PLATE 17
_A._ Cross-section of Mandrake Rhizome (_Podophyllum peltatum_, L.).
1. Epidermis.
2. Phellogen.
3. Cortical parenchyma.
_B._ Stone cell periderm of white cinnamon (_Canella alba_, Murr.).]
[Illustration: PLATE 18
PERIDERM OF WHITE OAK (_Quercus alba_, L.)
1. Outer layer of cork cells. 2. Cortical parenchyma cells. 3.
Stone cells. 4. Phellogen. 5. Cortical parenchyma cells.]
ORIGIN OF CORK CELLS
The cork cells are formed by the meristimatic phellogen cells, which
originate from cortical parenchyma. These cells divide into two
cells, the outer changing into a cork cell, while the inner cell
remains meristimatic. In other instances the outer cell remains
meristimatic, while the inner cell changes into a cortical parenchyma
cell. The development of a cortical parenchyma cell from a divided
phellogen cell is shown in Plate 101, Fig. 6. Both the primary and
secondary cork cells originate from the phellogen or cork cambrium
layer. Cork cells do not contain living-cell contents; in fact, in
the majority of medicinal barks the cork cells contain only air.
The walls of typical cork cells are composed, at least in part, of
suberin, a substance which is impervious to water and gases. In
certain cases layers of cellulose, lignin, and suberin have been
identified. Suberin, however, is present in all cork cells, and in
some cases all of the walls of cork cells are composed of suberin.
Suberized cork cells are colored yellow with strong sodium hydroxide
solutions and by chlorzinciodide.
CHAPTER III
MECHANICAL TISSUES
The =mechanical tissues= of the plant form the framework around
which the plant body is built up. These tissues are constructed and
placed in such a manner in the different organs of the plant as to
meet the mechanical needs of the organ. Many underground stems and
roots which are subjected to radial pressure have the hypodermal
and endodermal cells arranged in the form of a non-compressible
cylinder. Such an arrangement is seen in sarsaparilla root (Plate
38, Fig. 4). The mechanical tissue of the stem is arranged in the
form of solid or hollow columns in order to sustain the enormous
weight of the branches. In roots the mechanical tissue is combined
in ropelike strands, thereby effectively resisting pulling stresses.
The epidermis of leaves subjected to the tearing force of the wind
has epidermal cells with greatly thickened walls, particularly at the
margin of the leaf. The epidermal cells of most seeds have very thick
and lignified cell walls, which effectively resist crushing forces.
The cells forming mechanical tissues are: bast fibres, wood
fibres, collenchyma cells, stone cells, testa epidermal cells, and
hypodermal and endodermal cells of certain plants. The walls of the
cells forming mechanical tissues are thick and lignified, with the
exception of the collenchyma cells and a few of the fibres. Lignified
cells are as resistive to pulling and other stresses as similar sized
fragments of steel. The hardness of their wall and their resistance
to crushing explain the fact that they usually retain their form in
powdered drugs and foods.
BAST FIBRES
One of the most important characters to be kept in mind in studying
bast fibres is the structure of the wall. In fact, the author’s
classification of bast fibres is based largely on wall structure.
Such a classification is logical and accurate, because it is based
upon permanent characters. Another character used in classifying bast
fibres is the nature of the cell, whether branched or non-branched.
In fact, this latter character is used to separate all bast fibres
into two fundamental groups--namely, branched bast fibres and
non-branched bast fibres. The third important character utilized in
classifying fibres is the presence or absence of crystals.
Bast fibres are classified as follows:
1. =Crystal bearing.=
2. =Non-crystal bearing.=
The crystal-bearing fibres are divided into two classes:
1. =Of leaves.=
2. =Of barks.=
The non-crystal bearing are divided into:
1. =Branched.=
2. =Non-branched.=
The branched and non-branched are divided into four classes:
1. =Non-porous and non-striated.=
2. =Porous and non-striated.=
3. =Striated and non-porous.=
4. =Porous and striated.=
CRYSTAL-BEARING BAST FIBRES
The =crystal-bearing fibres= are composed (1) of groups of fibres,
(2) of crystal cells, and (3) of crystals. In these cases the groups
of fibres are large, and they are frequently completely covered by
crystal cells, which may or may not contain a crystal. The crystals
found on the fibres from the different plants vary considerably in
size and form. As a rule, the fibres when separated are free of
crystal cells and crystals. This is so because the crystal cells
are exterior to the fibres, and in separating the fibres during
the milling process the crystal cells are broken down and removed
from the fibres. It is common, therefore, to find isolated fibres
and crystals associated with the crystal-bearing fibres. The fibres
which are crystal-bearing may be striated or porous, etc.; but owing
to the fact that the grouping of the fibres and crystals is so
characteristic, little or no attention is paid to the structure of
the individual fibres.
[Illustration: PLATE 19
CRYSTAL-BEARING FIBRES OF BARKS
1. Frangula (_Rhamnus frangula_, L.).
2. Cascara sagrada (_Rhamnus purshiana_, D.C.).
3. Spanish licorice (_Glycyrrhiza glabra_, L.).
4. Witch-hazel bark (_Hamamelis virginiana_, L.).]
=Crystal-bearing fibres= occur in the barks of frangula (Plate 19,
Fig. 1); cascara sagrada (Plate 19, Fig. 2); witch-hazel (Plate 19,
Fig. 4); in cocillana (Plate 20, Fig. 1); in white oak (Plate 20,
Fig. 2); in quebracho (Plate 20, Fig. 3); and in Spanish licorice
root (Plate 19, Fig. 3).
The crystal-bearing fibres of leaves are always associated with
vessels or tracheids and with cells with chlorophyl. The presence
or absence of crystal-bearing fibres in leaves should always be
noted. The crystal-bearing fibres of leaves are composed of fragments
of conducting cells, fibres, crystal cells, and crystals. The
crystal-bearing fibres of leaves occur in larger fragments than the
other parts of the leaf, because the fibres are more resistant to
powdering. Having observed that a leaf has crystal-bearing fibres,
in order to identify the powder it is necessary to locate one of the
other diagnostic elements of the leaf--as the papillæ of coca (Plate
21, Fig. 1), or the hair of senna (Plate 21, Fig. 3), or the vessels
in eucalyptus (Plate 21, Fig. 2).
=Branched bast fibres= occur in only a few of the medicinal plants,
notable examples being tonga root and sassafras root. Occasionally
one is found in mezereum bark.
The bast fibre of tonga root (Plate 22, Fig. 2) often has seven
branches, but four- and five-branched forms are more common. The
walls are non-porous, non-striated, and nearly white.
The bast fibre of sassafras (Plate 22, Fig. 1) has thick, non-porous,
and non-striated walls, and the branching occurs usually at one end
only of the fibre. Most of the bast fibres of sassafras root are
non-branched.
POROUS AND STRIATED BAST FIBRES
=Porous and striated= walled bast fibres occur in blackberry bark of
root, wild-cherry bark, and in cinchona bark.
The fibres of blackberry root bark (Plate 23, Fig. 1) have distinctly
porous and striated walls; the cavity, which is usually greater than
the diameter of the wall, contains starch. These fibres usually occur
as fragments.
In wild-cherry bark (Plate 23, Fig. 2) the fibre has short, thick,
unequally thickened walls, which are porous and striated. Most of the
fibres are unbroken.
[Illustration: PLATE 20
CRYSTAL-BEARING FIBRES OF BARKS
1. Cocillana (_Guarea rusbyi_, [Britton] Rusby).
2. White oak (_Quercus alba_, L.)
3. Quebracho (_Aspidosperma quebracho-blanco_, Schlechtendal).]
[Illustration: PLATE 21
CRYSTAL-BEARING FIBRES OF LEAVES
1. Coca leaf (_Erythroxylon coca_, Lam.).
2. Eucalyptus leaf (_Eucalyptus globulus_, Labill).
3. Senna leaf (_Cassia angustifolia_, Vahl.).]
[Illustration: PLATE 22
BRANCHED BAST FIBRES
1. Sassafras root bark (_Sassafras variifolium_, [Salisb.] Kuntze).
2. Tonga root.]
Yellow cinchona bark (Plate 23, Fig. 3) has very thick, prominently
striated porous-walled fibres, with either blunt or pointed ends. The
cavity is narrow, and the pores are simple or branched.
POROUS AND NON-STRIATED BAST FIBRES
=Porous and non-striated= bast fibres occur in marshmallow root and
echinacea root.
The fibres of marshmallow (Plate 24, Fig. 3) usually occur in
fragments. The walls have simple pores, and the diameter of the
cell cavity is very wide; the pores on the upper or lower wall are
circular or oval in outline (end view).
The bast fibres of echinacea root (Plate 24, Fig. 4) are seldom
broken; the walls are yellow, the pores are simple and numerous. The
edges and surface of the fibres are frequently covered with a black
intercellular substance.
NON-POROUS AND STRIATED BAST FIBRES
=Non-porous and striated= bast fibres occur in elm bark, stillingia
root, and cundurango bark. The bast fibres of elm bark (Plate 25,
Fig. 1) occur in broken, curved, or twisted fragments. The central
cavity is very small, and the walls are longitudinally striated.
In powdered stillingia root (Plate 25, Fig. 2) the bast fibres are
broken, and the wall is very thick and longitudinally striated. The
central cavity is small and usually not visible. Bast fibres of
cundurango (Plate 25, Fig. 3) are broken in the powder. The cavity
is very narrow, and the striations are arranged spirally, less
frequently transversely.
NON-POROUS AND NON-STRIATED BAST FIBRES
=Non-porous and non-striated= walled bast fibres occur in mezereum
bark, in Ceylon cinnamon, in sassafras root bark, and in soap bark.
The simplest non-porous and non-striated walled bast fibres are found
in mezereum bark (Plate 26, Fig. 4). The individual fibre is very
long. It often measures over three millimeters in length, so that in
the powder the fibre is usually broken. The wall is non-lignified,
white, non-porous, and of uniform diameter.
[Illustration: PLATE 23
POROUS AND STRIATED BAST FIBRES
1. Blackberry root (_Rubus cuneifolius_, Pursh.).
2. Wild cherry (_Prunus serotina_, Ehrh.).
3. Yellow cinchona (_Cinchona species_).]
[Illustration: PLATE 24
POROUS AND NON-STRIATED BAST FIBRES
1. Sarsaparilla root (Hypoderm), (_Smilax officinalis_, Kunth).
2. Unicorn root (Endoderm).
3. Marshmallow root (_Althæa officinalis_, L.).
4. Echinacea root (_Echinacea angustifolia_, D. C.).]
[Illustration: PLATE 25
NON-POROUS AND STRIATED BAST FIBRES
1. Elm bark (_Ulmus fulva_, Michaux).
2. Stillingia root (_Stillingia sylvatica_, L.).
3. Cundurango root bark (_Marsdenia cundurango_, [Triana] Nichols).]
In Ceylon cinnamon (Plate 26, Fig. 2) the bast fibres measure up to
.900 mm. in length, so that in powdering the bark the fibre is rarely
broken. These bast fibres, unlike the bast fibres of mezereum, have
thick, white walls and a narrow cell cavity. Both ends of the fibre
taper gradually to a long, narrow point.
In Saigon cinnamon the bast fibres are not as numerous as they are
in Ceylon cinnamon. The individual fibres are thicker than in Ceylon
cinnamon, and the walls are yellowish and rough and the ends bluntly
pointed. These fibres are rarely ever free from adhering fragments of
parenchyma tissue.
In sassafras root bark (Plate 26, Fig. 3) the fibre has one nearly
straight side--the side in contact with the other bast fibres--and
an outer side with a wavy outline, caused by the fibre’s pressing
against parenchyma cells, the point of highest elevation being the
point of the fibre’s growth into the intercellular space between two
cells. The outer part of the wall tapers gradually at either end to a
sharp point. The walls are white, thick, and non-porous.
In soap bark (Plate 26, Fig. 1) the bast fibres have thick, white,
wavy walls and a narrow cavity. One end of the cell is frequently
somewhat blunt while the opposite end is slightly tapering.
The branched stone cells of wild-cherry bark have three or more
branches. The pores are small and usually non-branched, and the
striations are very fine and difficult to see unless the iris
diaphragm is nearly closed. The central cavity is very narrow and
frequently contains brown tannin.
The branched stone cells of hemlock bark are very large; the walls
are white and distinctly porous bordering on the cell cavity, which
contains bright reddish-brown masses of tannin.
In cross-section bast fibres occur singly or isolated, as in Saigon
cinnamon (Plate 34, Fig. 1); or in groups, as in menispermum (Plate
27, Figs. 1 and 2); or in the form of continuous bands, as in buchu
stem (Plate 100, Fig. 5).
Bast fibres are seen in longitudinal view in powdered drugs. The cell
cavity shows throughout the length of the fibre. This cavity differs
greatly in different fibres. In soap bark (Plate 26, Fig. 1) there is
scarcely any cell cavity, while in mezereum bark (Plate 26, Fig. 4)
the cell cavity is very large.
[Illustration: PLATE 26
NON-POROUS AND NON-STRIATED BAST FIBRES
1. Soap bark (_Quillaja saponaria_, Molina).
2. Ceylon cinnamon bark (_Cinnamomum zeylanicum_, Nees).
3. Sassafras root bark (_Sassafras variifolium_, [Salisb.] Kuntze).
4. Mezereum bark (_Daphne mezereum_, L.).]
[Illustration: PLATE 27
GROUPS OF BAST FIBRES
1. Menispermum rhizome (_Menispermum canadensis_, L.).
2. Althea root (_Althæa officinalis_, L.) showing two groups of bast
fibres.]
The pores, which are absent in many drugs, are, when present,
either simple, as in echinacea root (Plate 24, Fig. 4), or they are
branched, as in yellow cinchona (Plate 23, Fig. 3).
In each of the above fibres the length and width of the fibre
are shown. The fibres also have pores of variable length. Such a
variation is common to most fibres with pores. That part of the wall
immediately over or below the cell cavity shows the end view or
diameter of the pore, as in the fibre of marshmallow root (Plate 24,
Fig. 3). As a rule, however, the pores show indistinctly on the upper
and lower wall.
OCCURRENCE IN POWDERED DRUGS
In powdered drugs bast fibres occur singly or in groups. The
individual fibres may be broken, as in mezereum and elm bark, or they
may be entire, as in Ceylon cinnamon and in sassafras bark (Plate 26,
Figs. 2 and 3).
The lignified walls of bast fibres are colored red by a solution of
phlorogucin and hydrochloric acid, and the walls are stained yellow
by aniline chloride.
In fact, few of the fibres found in individual plants occur in a
broken condition.