LOCATION OF THE CAROTENOID PIGMENTS OF COEYNEBACTERIUM
SPECIES STRAIN 7E1C
APPROVED:
Major Professor
Minor7Professo
Director of the Department ^f Biology
Dean lof the Graduate School
/'//
Wilkinson, Joanne C., Location of the Carotenoid Pigments
of Corynebaaterium Species Strain 7E1C.
Master of Science
(Biology), August, 1973, 33 pp., 1 table, 4 figures, biblioggraphy, 76 titles.
Carotenoid pigments occur in a large number of red, orange
and yellow nonphotosynthetic microorganisms of the genera
Mycobacterium3
Nocardia and Micrococcus.
The chromogenic
organism tentatively named Corynebaaterium species strain
7E1C is closely related taxonomically to Nocardia and Mycobacterium.
This strain produces carotenoid pigments which
are xanthophylls rather than carotenes.
In most of the
organisms investigated so far, the carotenoids are found in
the cell membrane where they perform an unknown role, although
several functions have been hypothesized.
The purpose of this investigation was to determine the
site of the carotenoid pigments in C. spp. strain 7E1C as a
step towards resolving the role of the pigment in the cell.
Initially, protoplast formation of C. spp. strain 7E1C
was attempted by a number of procedures used successfully with
other microorganisms.
It was reasoned that production of pig-
mented protoplasts would indicate that the carotenoids were
located in the cell membrane or cytoplasm.
On the other hand,
pigmentless protoplasts would indicate that the cell wall was
the site of carotenoid pigments.
protoplasts were unsuccessful.
All attempts to produce
As an alternate approach to the determination of the
site of the carotenoids, a pure cell wall fraction was prepared by differential centrifugation and chemical treatment
of broken cells.
The purity of the cell wall preparation
was verified by electron microscopy.
Pigment assays of the individual fractions were estimated
1^
using
) of 2S00 at 483 nm.
6 an extinction coefficient vC E f
1 cnr
Approximately eighteen times as much pigment was located in
the cell membrane as in the cell wall.
The small amount of
pigment found in the cell wall may be due to trace cell membrane contamination of the cell wall.
Since function is
presumably coincident with location, carotenoid pigments of
C. spp. strain 7E1C may serve some purpose in stabilization or
protection at the membrane level.
Carotenoids of C. spp. strain 7E1C cannot be extracted
completely with the usual solvents but only when these solvents
are used in conjunction with a substance that specifically
solubilizes protein.
These results indicate that the pig-
ments may be bound in some way.
This linkage may be similar
to the carotenoid-glycosidic bond demonstrated in Sarcina
morrhuae and as such could serve as a means of stabilizing the
membrane.
LOCATION OF THE CAROTENOID PIGMENTS OF COEYNEBACTERIUM
SPECIES STRAIN 7E1C
THESIS
Presented to the Graduate Council of the
North Texas State University in Partial
Fulfillment of the Requirements
For the Degree of
MASTER OF SCIENCE
By
Joanne C. Wilkinson, B. S.
Denton, Texas
August, 1973
TABLE OF CONTENTS
Page
LIST OF TABLES
iv
LIST OF ILLUSTRATIONS
v
INTRODUCTION
1
MATERIALS AND METHODS
6
RESULTS AND DISCUSSION
11
CONCLUSION
24
SUMMARY
26
REFERENCES
27
in
LIST OF TABLES
Table
I.
Page
Pigment Recovery from Differential
Centrifugation
IV
19
LIST OF ILLUSTRATIONS
Figure
Page
1.
Purification of Cell Walls
2.
Whole Cells of Corynebaotevium
Strain 7E1C
3.
4.
13
Species
15
Cells of Corynebaoterium Species Strain
7E1C After Disruption in the MSK Cell
Homogenizer.
17
Purified. Cell Walls of Corynebacterium
Species Strain 7E1C
18
INTRODUCTION
"Carotenoids are a class of hydrocarbons (carotenes)
and their oxygenated derivatives (xanthophylls) consisting of
eight isoprenoid units joined in such a manner that the arrangement of the isoprenoid units is reversed at the center of the
molecules so that the two central methyl groups are in a 1,5
positional relationship" (25).
These compounds are chemically
related to lycopene, the tomato pigment, and are widely distributed in nature (74).
In photosynthetic organisms,
carotenoids participate in light absorption for photosynthesis,
protection of chlorophylls from light destruction and in some
way contribute to the complex infrared spectrum of bacteriochlorophyll (12,28,59,74).
Several fungi (10,20,21,26,27)
and many yellow, red and orange pigmented nonphotosynthetic
bacteria (1,4,11,30,31,34,37,42,60,63,65,63) also contain
carotenoid pigments.
The presence of carotenoid pigments in
microorganisms seems to have taxonomic significance only in
the algae (10).
An integral part in the determination of the relationship of carotenoid pigments of nonphotosynthetic bacteria to
cell functions is the localization of the carotenoids in the
cell.
In Savoina lutea, for example, the carotenoid pigments
are located in the cell membrane (45) and are thought to
prevent photodynamic killing by either internal or external
photosensitizers (44) . Micrococcus lyeodeikticus (54),
Sarcina flava (66), Staphylococcus aureus (31), Halobacterium
halobium and Halobacterium salinarium (61) also contain
carotenoids which seem to be associated with the cell membrane
rather than with the cell wall.
On the other hand, carotenoids
are located in the cell wall of Micrococcus radiodurans (76)
and in the "knallgas" bacterium 12/60/x (15) .
Work and
Griffith (76) showed that the site of carotenoid pigments
was the middle layer of the three-layered cell wall of M.
radiodurans.
Salton and Freer (55) considered carotenoid pigments to
be localized almost exclusively (at least 951) in the cell
membrane of microorganisms and suggested that carotenoids
could be used as "marker molecules" in the fractionation of
these membranes.
Salton and Ehtisham-ud-Din (54) postulated
that, although the carotenoids are not indispensable, they
contribute to membrane stability in S. lutea and M. lysodeikticus.
Additional theories of carotenoid function in the
bacterial cell include protection from oxidizing agents (26)
and protection from photo-oxidation by visible light (37,43).
Studies utilizing Mycobacterium marinum showed that the
efficiency of photoprotection was directly proportional to
the amount of carotenoid pigment per cell (42) .
Since carot-
enoids are generally not produced in cultures maintained in
the dark but are produced in response to light stimulus (3,63),
a photoprotective function may be a valid hypothesis for these
organisms.
Formation of a spheroplast or protoplast of a chromogenic
organism is useful in the determination of the site of pigment
location (45).
The term "protoplast" is used to denote the
structure remaining when the cell wall is removed from the
bacterial cell (48).
"Spheroplasts" are osmotically fragile,
spherical cells but still retain part of the cell wall (48).
If the pigments are released into the media when a protoplast
is formed, presumably the pigments are localized in the cell
wall.
On the other hand, if protoplast formation is complete
and all cell wall material is removed, but the cells en masse
are pigmented, then the carotenoids are said to be in the
cell membrane or the cytoplasm.
Lysis of the protoplast by
osmotic shock can then be used to determine the precise
location of the pigments.
Protoplasts can be formed by a variety of methods.
One
of the most commonly used agents for the production of protoplasts is lysozyme which causes the lysis of many gram positive
bacteria.
Lysozyme catalyzes the hydrolysis of the l-*-4 gly-
cosidic linkages in the polymer peptidoglycan, N-acetyl
glucosamine —N-acetyl muramic acid (57) .
Pencillin has also
been used for protoplast production, especially with gram
negative microorganisms (40) .
This antibiotic inhibits the
terminal transpeptidation reaction in which the linear peptidoglycan becomes cross-linked to another strand (62).
Once
this linkage is established it cannot be reversed by penicillin.
By way of contrast, the mode of protoplast formation by
D-cycloserine is, theroretically, interference with the biosynthesis of the uredine nucleotide precursors (62) .
Glycine
induction of protoplasts is presumably due to reversal of the
terminal crosslinking reaction in cell wall synthesis (62).
Enzymes that solubilize lipid (16) or protein outer layers of
the wall (1,36), or chelating agents (48) that bind ions
essential for the synthesis of the cell wall and for its
stability, have also been used in conjunction with lysozyme,
glycine or penicillin to produce protoplasts.
In organisms that do not form protoplasts, disruption
of bacterial cells by mechanical means and preparation of a
pure cell wall fraction by differential centrifugation has
been used as a means of determining the location of carotenoids (45) . Assay of total pigment at every step of the
process is necessary for determining the cell fraction in
which the pigment is located.
Corynebacterium spp. strain 7E1C has a cell wall composition similar to those of Mycobacterium species and
Nocardia species.
All of these genera have a high content
of lipid in the cell wall.
Preparation of pure cell walls of
these organisms is difficult because the lipid apparently
enhances entrapment of cell membrane fragments (53) . According
to Salton (53), electron microscopy is the best single method
of validating the purity of cell wall material.
The taxonomic position of the organism tentatively named
Corynebaaterium species strain 7E1C (33), one of the "strains
in search of a genus" (19), is still uncertain (5,30,39,41,68,
72,73).
This strain is a chromogenic organism whose photo-
inducible carotenoid pigments have been partially characterized.
The carotenoid pigments are xanthophylls rather than carotenes
(30); six hypophasic and three epiphasic pigments have been
detected so far.
Cardini and Jurtshuk (9) have reported
difficulty with contamination of enzymatically active fractions,
presumably of membrane origin, of C. spp. strain 7E1C with a
carotenoid pigment that absorbs at 450 nm.
Their investigation
did not determine whether the carotenoid is located in the cell
membrane, or is an absorbed contaminant.
Also, they did not
report whether all of the carotenoid or just one of the nine
pigments is the cause of their difficulty.
This investigation attempts to show the location of the
carotenoid as a step towards resolving the role of the pigment
in the cell but makes no attempt to prove the function of the
carotenoid pigments to Corynebacterium spp. strain 7E1C.
MATERIALS AND METHODS
The organism used in this investigation, Corynebacterium
species strain 7E1C, was originally isolated from soil by
Kester and Foster (33) on mineral salts media enriched with
propane as the sole carbon and energy source.
The organism
was grown in "L-salts" media, according to the formula of
Leadbetter (38), supplemented with 1% dextrose.
Flasks were
incubated at room temperature (22-25 C) on an Eberbach rotary
shaker at 140 revolutions per minute at a rotation diameter of
1.2 inches.
Two per cent agar was added to the L-salts medium
for slants on which stock cultures were maintained.
Inocula
for liquid cultures were prepared from logarithmic phase cells
grown in L-salts media.
The cell density was adjusted to an
optical density of 0.30 at 580 nm by dilution with sterile
L-salts.
One part of this suspension was added to 99 parts
of L-salts media.
Purity of inoculum was determined by streak
plates on L-salts agar and by microscopic examination.
Protoplast Production —Procedures used for protoplast
formation were those of
Lederberg (40) (penicillin G), Jeynes
(29) (glycine), Hines et al. (23) (glycine and penicillin),
Gooder (17) (lysozyme), Heppel (22) (lysozyme and ethylenediaminetetraacetic acid (EDTA)), Noller and Hartsell (49)
(lysozyme, trypsin and butanol), Warren et al. (71) (polymyxin),
Kohn (36) (freeze-thaw and lysozyme), Barker and Thorne (2)
(trypsin, lysozyme and EDTA), Ghosh and Murray (16) (lipase,
bile salts and lysozyme) and Mattman (46) (D-cycloserine).
Since C. spp. strain 7E1C has a long generation time,
organisms for protoplast production were grown in L-salts
media for four days at room temperature at which time they
were in early logarithmic phase.
Organisms were incubated
for six days, at which time plate counts yielded about 1.2 x
Q
10
cells per ml, for techniques requiring stationary phase
cultures.
Cells were grown in the presence of 1 mM magnesium
chloride in experiments utilizing EDTA for protoplast formation, since magnesium ion limitation negates the effect of
EDTA.
Cells were examined by phase-contrast microscopy for
protoplast formation.
They were also checked for osmotic
fragility by dilution of treated and untreated samples in
equal volumes of either distilled water or 1.41 M sucrose.
The dilutions were measured spectrophotometrically and any
decrease in optical density was recorded.
Cell Wall Preparation —Cells used as a source of pure
cell walls were grown in L-salts for five days to yield a
late logarithmic phase culture.
The organisms were collected
by continuous flow centrifugation at 23,500 x g in a Sorvall
Model SS-3 centrifuge.
After one wash in 0.85% NaCl (pH 7.0),
the cells were centrifuged and the sedimented cells resuspended
in 0.851 NaCl to yield a sediment with the consistency of
heavy cream.
In order to facilitate disruption, this bac-
terial suspension was frozen at -20 C and then thawed to room
temperature in warm water.
Glass beads of 0.10 to 0.11 mm in
diameter were added to give a ratio of one part beads to five
parts cell suspension.
The bacteria-bead mixture was agitated
in an MSK cell homogenizer at 2000 rpm for two minutes under a
constant stream of liquid carbon dioxide to prevent excessive
heating.
The suspension was observed by phase-contrast micro-
scopy to ascertain the degree of breakage.
When no whole cells
could be observed, the broken cells were subjected to differential centrifugation in a Sorvall Model SS-3 centrifuge
according to the method of Takeya and Hisatsune (64).
The
crude cell wall material was washed once with 2% Triton X-100
as recommended by Schnaitman (57,58).
Specimens from each step
of the differential centrifugation procedure were lyophilized
for subsequent preparation for electron microscopy.
Assay of Total Pigment —Carotenoid pigments were obtained
from each fraction by one extraction with a 1:5 (v/v) solution
of carbon disulfide in methanol followed by six subsequent
extractions with carbon disulfide alone.
Since carbon disul-
fide has a density greater than that of the whole cells and
cell fragments, the utilization of carbon disulfide was somewhat more inconvenient.
However, it was the preferred solvent
because it gave more complete extraction of total cell pigments
The pigment extracts were pooled and evaporated to dryness at
40 C in a Buchler flash evaporator.
In order to remove residual
carbon disulfide and methanol, the dried pigment was solubilized
in chloroform, re-evaporated and finally taken up in a small
amount of chloroform.
Estimation of the total pigment in a sample was obtained
1 9by using an extinction coefficient ( E ^ ) of 2500 at the wavelength of maximum absorption, 483 nm in chloroform.
If "X"
grams of carotenoid dissolved in "y" nil of solvent give an
extinction coefficient of "E" at its wavelength of maximum
absorption, then:
Ey
X
=
(13).
E
1%
x
100
1 cm
Spectrophotometry assays were performed on a Bausch and Lomb
Spectronic 20.
Electron Microscopy —Specimens for electron microscopy
were fixed by a method similar to that of Cagle et al. (8).
Initial fixation was performed in 0.151 ruthenium red (w/v)
in 0.1 M sodium cacodylate buffer (pH 7.2) for thirty minutes
followed by fixation in 41 glutaraldehyde in 0.133 M cacodylate buffer and 0.05% (w/v) ruthenium red for one hour.
After
one wash in cacodylate buffer, the specimens were further fixed
in 1.331 osmium tetraoxide and 0.051 (w/v) ruthenium red in
0.133 M cacodylate buffer for one and one-half hours.
A final
10
wash in 0.2 M cacodylate buffer was followed by dehydration in
ethyl alcohol.
The dehydrated specimens were embedded in Epon
812 and polymerized at 60 C for 24-48 hours.
Thin sections
were cut with a Sorvall Porter-Blum MT-2 ultramicrotome with
glass knives.
The thin sections were stained with a saturated
uranyl acetate stain in 30% ethanol for three minutes and then
with lead citrate (47) for five minutes.
After carbon coating
(32), the sections were examined in an RCA EMU-3G electron
microscope at an accelerating voltage of 50 kv.
Photographs
were taken on Kodak 2 inch by 10 inch projector slide plates
of medium contrast and fine grain film.
RESULTS AND DISCUSSION
Initially, protoplast
production was considered neces-
sary in order to determine the location of the carotenoid
pigments.
It was reasoned that if a pigmented protoplast
resulted from these preparations, the site of pigment would
be the cell membrane or the cytoplasm.
A nonpigmented proto-
plast would indicate that the cell wall was the location of the
carotenoids.
Preparations of protoplasts were attempted by
using a broad range of concentrations of penicillin (40),
glycine (23,29,51), lysozyme (17), EDTA (22), D-cycloserine
(46) and polymyxin (71) . Freezing and thawing (36) and combinations of these chemicals (2,22,23,49) was also investigated.
In addition, the effect of temperature on conversion of cells
to protoplasts by the substances given above was determined.
Stabilization of protoplasts was attempted with sucrose concentrations ranging from 0.25 M-2.0 M.
Production of proto-
plasts by any of these methods or any adaptations of these
methods was unsuccessful.
In light of similar unsuccessful
attempts with related organisms (48), this failure was not
unexpected.
Methods utilizing glycine addition to logarithmic
phase cultures in concentrations of 2% in 0.6 M sucrose did
produce spherical cells rather than the typical short bacillus,
but these were not osmotically fragile and thus could not be
considered to be protoplasts.
11
Penicillin concentrations of
12
100 units per ml resulted in aggregation of the cells upon
standing but did not alter the shape of the organism even
after incubation for prolonged periods of time.
The failure
of lysozyme to effect the cell wall may well be due to a
permeability factor as well as to the hydrophobic nature of
the organism.
The waxy outer layer of the organism may
prevent chemicals from reacting with the wall structure.
All
attempts to remove the wax layer with detergents, solvents
and enzymes prior to treatment were unsuccessful.
Cell Rupture and Differential Centrifugation —
Due to
failure to produce protoplasts, it was necessary to attempt
an alternate approach.
Rupture of bacterial cells followed
by differential centrifugation is useful in determining the
location of the pigments (45) .
Freezing and thawing the cells
just prior to disruption was necessary for efficient cell
rupture.
Plate counts made of the bacterial suspensions imme-
diately before and after rupture revealed a 99.991 reduction
in viable cells.
The disrupted suspensions were observed by
phase-contrast microscopy as small amorphous aggregates of
material not recognizable as intact bacterial cells.
The pH
of the disrupted suspensions was also checked since some glass
beads release alkali when shaken.
No observable pH change was
noted.
Purification of the cell wall was performed according to
the scheme outlined in Figure 1.
Trypsin, 0.2% in 0.067 M
13
FIGURE 1
PURIFICATION OF CELL WALLS
Whole Cells
rupture; coarse filter
on a sintered glass filter;
centrifuge at 2300 x G, 10
minutes, twice
1
Supernatant I
Sediment I
coarse debris and unbroken
cells; wash once with 0.851
NaCl and add wash to
Supernatant I
Sediment IB
centrifuge at 25,000 x G, 15 minutes
I
Supernatant II
Sediment II
cell membrane
cell wall and cell
membrane
j
wash once in 1 M NaCl; treat
with 0.2% trypsin at 37 C for
3 hours; centrifuge at 25,000
x G, 10 minutes
j
Sediment III
Supernatant III
crude cell wall
cell membrane fragments
Supernatant IV
Supernatant of washes
-wash
wash
wash
wash
I
#1; phosphate buffer
#2; 21 Triton X-100
#3-#5; 1 M NaCl
#6-#8; distilled water
Sediment IV
pure cell wall
14
phosphate buffer (pH 8.0), was incubated with the cell wallcell membrane fraction for three hours at 37 C to selectively
solubilize the proteins of the cell membrane.
Trypsin treat-
ment has no effect on the cell walls of gram positive bacteria
(64).
After subsequent washing with phosphate buffer, the
crude cell wall fraction was treated with 2% Triton X-100 in
phosphate buffer for twenty minutes at 24 C.
Triton X-100
specifically solubilizes proteins of the cell membrane (14,57,
58).
This was followed by thorough washing with 1 M NaCl and
distilled water to wash away the dissolved cell membrane
material.
The remaining sediment, which was virtually non-
pigmented, was verified by electron microscopy as consisting
entirely of cell wall with little or no discernable cell
membrane contamination.
Figure 2 is an electron microscope
photograph of whole cells of C. spp. strain 7E1C.
Two or more
globose, lipid-like inclusion bodies are readily observed in
this cell.
These inclusion bodies are reminiscent of the
"sap-vacuoles" (2), of unknown substances, evident in thin
sections of Mycobacterium tuberculosis var. avium and a
variant of Mycobacterium -phlei. The small, irregularly shaped
bacillus seen in Figure 2 also demonstrates an intact cell
with cell wall and cell membrane continuous around the cell.
One mesosome-like organelle can be seen as an elaboration of
the membrane in the lower portion of the cell.
Inclusion
bodies resembling polyphosphate can also be noted as round
electron-dense areas in the cytoplasm.
The irregular edge
15
*
jk-
Fig. 2. Whole cells of Corynebaoterium
species strain 7E1C
The cell wall (CW) is separated from the cell membrane (CM)
by a periplasmic space (PS) and seems to be fairly typical
of walls normally surrounding gram-positive cells. A mesosome (M) is seen in the lower portion of the irregularlyshaped bacillus. Several lipid-like (L) sacs can also be
seen in this cell. Magnification is 161,350 x.
16
of the cell wall may well be due to the wax that is formed by
these organisms.
The appearance of cells after disruption in the MSK cell
homogenizer for two minutes is shown in Figure 3.
Micelles
of membrane, as well as membrane fragments and extraneous
cytoplasmic material absorbed to the cell wall, can be seen
in the photograph.
The gross morphology of the broken cells
is greatly different from that of the intact cell; no lipidlike sacs are observed intact and the cells have completely
lost their anatomic integrity.
The broken cells appear as
empty shells with the cell wall relatively intact.
These
cell fragments appear to be contaminated with extraneous
material.
The pure wall fraction (shown in Figure 4) demon-
strates the appearance of the broken cells after treatment
with trypsin, Triton X-100 and adequate washing and is in
marked contrast to the cell fragments seen in Figure 3.
The
pure cell wall fraction is free of cell membrane and cytoplasmic impurities but is itself relatively intact.
Thus,
the procedure used resulted in a pure cell wall preparation.
As shown in Table I, the majority of the pigment is in
the cell membrane fraction.
In fact, 100-1421 of the total
pigment accounted for in the Supernatant I was solubilized
with the membrane.
Oddly enough, a net yield of 111-145% of
the extractable pigment in Supernatant I (the broken cell
fraction) was obtained from the extraction of the individual
fractions.
The range of pigment values is used to indicate
17
Fig. 3. Cells of Corynebacterium species strain 7E1C after
disruption in the MSK cell homogenizer. Relatively intact
cell walls (CW) have adsorbed cytoplasmic material and cell
membrane (CM). Magnification is 66,800 x.
18
Fig. 4. Purified cell walls of Corynebactevium species strain
7E1C. Cell walls are apparently uncontaminated with intact
cell membrane. Magnification is 37,340 x.
19
TABLE I
PIGMENT RECOVERY FROM DIFFERENTIAL CENTRIFUGATION
Total Carotenoid
(10" 3 grams)
Fraction
Experiment
I
II
Percentage of
Original(6)
Experiment
I
II
Supernatant I
0.53
0.39
100
100
loss to E-M
0.06
0.03
11
7
Supernatant II(1)
0.33
0.01
62
0
Supernatant IIIC2)
0.05
0.02
9
4
Supernatant IVC3)
0.04
0.07
8
18
Sediment IV(4)
0.06
0.01
11
3
Supernatant of
washes
0.11
0.48
21
120
Carotenoid
attributable to
membrane(5)
0.53
0.57
100
142
1. cytoplasmic material and cell membrane
2. cell membrane fragments
3. cell membrane
4. pure cell wall
5. Supernatants II, III, IV and Washes
6. total extractable pigment in Supernatant I
20
the differences in recovery of pigment in repeated experiments
More complete membrane removal can be accomplished through
exhaustive washing.
However,this involved hours of washing
and centrifugation and is not significant when the additional
8-34%
pigment is compared to a total pigment yield of 111-
145%.
Original experimental procedures (64) required high centrifugation speeds.
These resulted in a packed cell button
which was difficult to resuspend.
Apparently, sufficient
agitation to resuspend the sediment resulted in dissolution
of the cell membrane.
Modified procedures were subsequently
followed with reduced, but adequate, centrifugation speeds as
listed in Figure 1.
In these procedures, washing was required
to release the cell membrane.
This demonstrates that the
carotenoid is not in the cytoplasm, as it appeared to be in
the original experiments, but rather in the cell membrane.
The reason for pigment recovery levels of 111-145% is
explained by the fact that cells and cell fragments that were
not treated with trypsin and Triton X-100 could not be completely extracted with carbon disulfide alone nor with carbon
disulfide and methanol.
Thus, the initial pigment concen-
trations, which was the basis for the calculation, was low.
Trypsin did not extract pigment from whole, unbroken cells.
When whole cells of C. spp. strain 7E1C were washed with 0.5%
Triton X-100 for one minute and then centrifuged, a deep
orange-red supernatant was recovered and the cell button was
21
pale pink.
According to Schnaitman (58), Triton X-100 specif-
ically solubilizes proteins of the cell membrane and has no
effect on the cell wall (57,58).
DePamphilis and Adler (14)
similarly reported that it does not have an effect on the
L-membrane.
Schnaitman further stated (58) that Triton X-100
is, in fact, specific for that portion of the cell membrane
that is closely bound to the cell wall.
Apparently, Triton
X-100 can penetrate the porous cell wall, solubilize the cell
membrane and release the carotenoids.
The pure cell wall fraction, after one extraction with
carbon disulfide and methanol, released all of its pigment
into the extract.
The broken, untreated cells still retained
a pink color even after ten extractions with carbon disulfide
over a period of seven hours.
This suggests that a small
portion of the pigment is bound in such a way, possibly a
carotenoid-protein linkage, that cannot be broken by solvents
but only by reagents that specifically break the bond.
Saperstein and Starr (56) reported that carotenoid pigments
of Corynebacteriat Mycobacteria and Micrococci occur as particulate protein complexes in the cell.
They hypothesized that
the pigment was necessary for the synthesis of the associated
protein.
They further stated that conditions which normally
cause isomerization of carotenoids when in solutions of organic
solvents do not affect carotenoids associated with protein,
possibly because of prevention of rotation around ethylene
bonds as a result of protein attachment (56).
A carotenoid
22
that was glycosidically bound to a glucose-bound peptide was
reported by Thirkell and Hunter in Saroina morrhuae (67) .
They reported failure in extracting all of the pigment with
solvents and later hypothesized that the bonding had to be
covalent to account for these results.
Synthetic detergents
freed these carotenoids from a membrane preparation.
A carot-
enoid-protein complex was identified by Howes and Batra (24)
as the photoreceptor in an unidentified Mycobacterium species.
The data thus far provided strongly infer the existence
of a carotenoid-protein moiety in C. spp strain 7E1C.
However,
although the evidence is highly suggestive, its existence can
be confirmed only by isolation and characterization of the
complex.
The distribution of the pigment as determined by assay
of the various purification steps is seen in Table I.
Only
3-121 of the total carotenoid can be accounted for by the pure
cell wall.
This residual carotenoid may be due to a minority
of cell membrane contamination or to absorption of a small
amount of carotenoid by the cell wall.
The cell wall may
serve as a reservoir for superflous carotenoid; excess produced
in the membrane may be deposited in the cell wall.
It is also
possible that the cell wall is the production or storage site
of one of the nine carotenoid pigments.
Even if a portion of
the pigment is found in the cell wall and is not a comtaminant,
the cell membrane still contains approximately eighteen times
as much carotenoid pigment as the cell wall.
The carotenoids
23
are most likely situated at the cell site where they function.
The location of the carotenoids on or in the membrane, points
to a role at that level.
Whether the role is one of pro-
tection from light or oxidizing agents, an alternate pathway
for electron transport in times of overproduction or a stabilizing agent to the membrane structure remains to be determined
CONCLUSION
Liquid cultures of C. spp. strain 7E1C grown in L-salts
media under fluorescent lighting at room temperature accumulate xanthophylls (30) .
An attempt to localize the pink-
orange pigment by formation of protoplasts was unsuccessful
since protoplasts could not be formed using known methods of
production.
Failure to produce a protoplast may be due to
the impermeability of the wax outer layer of the cell wall to
the usual agents employed in protoplast formation or to an
unusual cell wall structure.
As an alternate approach to the
determination of the pigment site in the cell, the organism
was efficiently disrupted in an MSK cell homogenizer and
purified cell walls prepared by differential centrifugation
according to the procedure of Takeya and Hisatsune (64) with
the modification of Schnaitman (58) .
Pigment analysis of
fractions obtained by differential centrifugation revealed
that the majority (in fact, nearly all) of the carotenoid was
found in the cell membrane, rather than in the cell wall or
free in the cytoplasm.
The small amount of carotenoid found
in the cell wall may be due to contamination with cytoplasmic
membrane or else the cell wall may indeed be a unique site for
a portion of the carotenoid.
Since location is usually coin-
cident with function, the carotenoid may serve some purposes
in protection or stability at the membrane level.
24
25
Inability to extract the entire carotenoid pigments from
broken cells that have not been treated with substances which
solubilize protein indicates that a portion of the carotenoid
in the cell may be bound in some fashion.
This linkage may
be similar to the carotenoid-glycosidic bond demonstrated in
S. morrhuae and as such could serve as a means of stabilizing
the membrane.
SUMMARY
Carotenoid pigments occur in a large number of red,
orange and yellow nonphotosynthetic bacteria of the genera
Mycobacteriumj Corynebaater-Lum and Nocardia.
In most of the
organisms investigated so far, the carotenoids are found in
the cell membrane where they perform an unknown role, although
several functions have been hypothesized.
Corynebaatevium spp.
strain 7E1C produces carotenoids which are xanthophylls (30)
rather than carotenes.
The majority of these pigments have
been found to be located in the cell membrane.
A small portion
of the total carotenoids cannot be extracted with solvents but
can be extracted completely from broken cell fragments after
treatment with chemicals that solubilize protein.
These
carotenoids may be similar to those of S. morrhuae in which a
carotenoid-glycoside peptide bond has been found.
Attempts to produce protoplasts of C. spp. strain 7E1C
by many procedures used successfully with other microorganisms
were not successful.
Differential centrifugation of broken
cells of the organism yielded a pure cell wall fraction uncontaminated with cytoplasmic membrane, as determined by electron
microscopic examination.
26
REFERENCES
1.
Aasen, A.J. and S. Liaaen Jensen. 1966. Carotenoids
of Flexibacteria:
IV. The carotenoids of two
further pigment types. Acta. Chem. Scand. 20:
2322-2324.
2.
Barker, D.C. and Kareen J.I. Thorne. 1970. Spheroplasts
of Lactobacillus casei and the cellular distribution of bactoprenol. J. Cell Sci. 7_: 755-757 .
3.
Batra, P.P. and H.C. Rilling. 1964. On the mechanism
of photoinduced carotenoid synthesis: Aspects of
the photoinductive reaction. Arch. Biochem. and
Biophys. 107 :485-492.
4.
Babkova, T.S. 1965. Carotenoid pigments of Mycobacteria
and yeasts. Microbiology. 34 : 229-233 .
5.
Bradley, S. Gaylen. 1971. Criteria for definition of
Mycobacteriums Nocardia and the rhodochrous complex.
Advan. Front, of Plant Sci. 28_:349-362.
6.
Brieger, E.M. 1963.
microorganisms.
7.
Bulgakova, V.G. and A.B. Silaev. 1971. Lytic effects of
certain polypeptide antibiotics on bacterial protoplasts (spheroplasts). Microbiology. 4£:92-95.
Structure and ultrastructure of
Academic Press. New York.
8.
Cagle, G.D., R.M. Pfister, and G.R. Vela. 1972. Improved
staining of extracellular polymer for electron
microscopy: Examination of Azotobactert Zoogloeas
Leuconostoc and Bacillus. Appl. Microbiol. 24:
477-488.
—
9.
Cardini, G. and Peter Jurtshuk. 1970. The enzymatic
hydroxylation of n-octane by Corynebacterium spp.
strain 7E1C. J. Biol. Chem. 245:2789-2796.
10.
Ciegler, Alex. 1965. Microbial carotenogenesis.
Advan. Appl. Micro. 7_:l-34.
11.
Courington, Doris P. and T.W. Goodwin. 1955. A survey
of the pigments of a number of chromogenic marine
bacteria, with special reference to the carotenoids.
J. Bacteriol. 70:568-571.
27
i
I
l
j
|
'
;
28
12.
Crouse, J.B., W.R. Sistrom, and S. Nemser. 1963.
Carotenoid pigments in the spectrum of bacteriochlorophyll. Photochem. Photobiol. 2^:361-375.
13.
Davis, B.H. 1965. Analysis of carotenoid pigments,
p. 489-531. In. T.W. Goodwin, Chemistry and biochemistry of plant pigments. Academic Press. New
York.
14.
DePamphilis, M.L. and Julius Adler. 1971. Purification
of intact flagella from Escherichia coli and
Bacillus subtilis. J. Bacteriol. 105:378 .
15.
Eberhardt, U. 1971. The cell wall as the site of
carotenoid in the "Knallgas" bacterium, 12/60/x.
Archiv. fur Mikrobiol. 80 : 32-37 .
16.
Ghosh, B.K. and R.G.E. Murray. 1967. Fine structure of
Listeria monocytogenes in relation to protoplast
formation. J. Bacteriol. 93:411-426 .
17.
Gooder, Harry. 1968. Protoplasts and L-forms of
Streptococcus. p. 42-43. Ill L.B. Guze (ed.),
Microbial protoplasts, spheroplasts and L-forms.
Th6 Williams and Wilkins Co. Baltimore, Maryland.
18.
Gordon, Ruth E. and Joan M. Mihm. 1959. A comparison
of four species of Mycobacterium. J. Gen. Microbiol.
21:736-748.
19.
Gordon, R.E. 1965. Some strains in search of a genusCorynebacterium3 Mycobacterium3 Nocardia, or
what? J. Gen. Microbiol. 43:329-343.
20.
Grebanovski-Sassa, Olga and F.H. Fappen. 1968. Light
and temperature effect of Epicoccum nigrum.
Phytochem. 7_: 1605-1617 .
21.
Haxo, Francis. 1948. Studies of the carotenoid pigments
of Neurospora: I. Composition of the pigment.
Arch. Biochem. 20:400-421.
22.
Heppel, Leon A. 1967. Selective release of enzymes
from bacteria. Science. 156:1451-1455 .
23.
Hines, William D., Bob A. Freeman, and Gary R. Pearson.
1964. Production and characterization of Brucella
spheroplasts. J. Bacteriol. 87:438-444.
29
24.
Howes, C.D. and P.P. Batra. 1970. Mechanism of
photoinduced carotenoid synthesis: Further
studies on the action spectrum and other aspects
of carotenogenesis. Arch. Biochem. and Biophys.
137:175-180.
25.
IUPAC commission on the nomenclature of organic chemistry
and IUPAC-IUB commission on biochemical nomenclature:
Tentative rules for the nomenclature of carotenoids.
1971. Biochem. 10:4827-4837.
26.
Jensen, Synn^ve Liaaen. 1965. Biosynthesis and function
of carotenoid pigments in microorganisms. Ann. Rev.
Microbiol. 19:163-172.
27.
Jensen, S. Liaaen and A.G. Andrews. 1972. Microbial
carotenoids. Ann. Rev. Microbiol. 26 : 225-248 .
28.
Jensen, Synn^ve Liaaen, Germaine Cohen-Bazere, T.O.M.
Nakayama, and R.Y. Stanier. 1958. The path of
carotenoid synthesis in a photosynthetic bacterium.
Biochim. Biophys. Acta. 29:477-498.
29.
Jeynes, M.H. 1957. Growth and properties of bacterial
protoplasts. Nature. 180 :867 .
30.
Jorgensen, Joyce A. 1971. Isolation and partial characterization of carotenoid pigments of Mycobacterium
rhodoahroue, unpublished Master's thesis, Department
of Biology, North Texas State University, Denton,
Texas.
31.
Joyce, Gay H. and David C. White. 1971. Effects of
benzo-(a)-pyrene and piperonyl butoxide on formation
of respiratory system phospholipids and carotenoids
of Staphylococcus avueus . J. Bacteriol. 106:403441.
32.
Juniper, B.E., A.J. Gilchrist, G.C. Cox, and P.R. Williams.
1970. Techniques for plant electron microscopy.
Blackwell Scientific Publications. Oxford, England.
33.
Kester, A.S. and J.W. Foster. 1962. Diterminal oxidation
of long-chain alkanes by bacteria. J. Bacteriol.
£5:860-869.
34.
Kleining, Hans, Hans Reichenbach, Hans Achenbach, and
Jochen Stadler. 1971. Carotenoid pigments of
Sorangium compositum (Myxobactevales) including
two new carotenoid glucoside esters and two new
carotenoid rhamnosides. Arch. Mikrobiol. 78:
224-233.
—
30
35.
Klieneberger-Nobel, E. 1960. L-forms of bacteria. Iji
I.C. Gunsalus and Roger Y. Stanier, The bacteria.
Volume I. p. 361-383. Academic Press. New York.
36.
Kohn, A. 1960. Lysis of frozen and thawed cells of
Escherichia coli by lysozyme, and their conversion
into spheroplasts. J. Bacterid. 79^:697-706.
37.
Kunisawa, Riyo and R.Y. Stanier. 1958. Studies on the
role of carotenoid pigments in a chemoheterotrophic bacterium, Corynebacterium -poinsettiae.
Archiv. fur Mikrobiol. 31:146-156 .
38.
Leadbetter, E.R. and J.W. Foster. 1958. Studies on
some methane-utilizing bacteria. Arch. Microbiol.
30:91-92.
39.
Lechevalier, Mary, Ann Horans, and H. Lechevalier. 1971.
Lipid composition in the classification of Nocardia
and Mycobacterium. J. Bacterid. 105 :313-318 .
40.
Lederberg, Joshua. 1956. Bacterial protoplasts induced
by penicillin. Proc. Nat. Acad. Sci. USA. 42:574577.
—
41.
Manion, R.E., S.G. Bradley, H.H. Zinneman, and W.H. Hall.
1964. Interrelationships between Mycobacterium and
Nocardiae. J. Bacterid. 87 :1056-1059 .
42.
Mathews, M. 1963. Localization, function, and formation
of the carotenoid pigments of a strain of Mycobacterium
marinum. Photochem. Photobiol. 2_:1-18.
43.
Mathews, Micheline M. and W.R. Sistrom. 1959. Function
of carotenoid pigments in non-photosynthetic bacteria.
Nature. 184:1892-1893 .
44.
Mathews, Micheline M. and W.R. Sistrom. 1960. The
function of carotenoid pigments of Sarcina lutea.
Archiv. fur Mikrobiol. 35 :139-146 .
45.
Mathews, Micheline M. and W.R. Sistrom. 1959. Intracellular location of carotenoid pigments of some
respiratory enzymes in Sarcina lutea. J. Bacteriol.
78_:778-787.
46.
Mattman, Lida H. 1968. L-forms isolated from infections.
p. 473. In Lucien B. Guze (ed.), Microbial protoplasts, spKeroplasts, and L-forms. The Williams and
Wilkins Co., Baltimore, Maryland.
31
47.
Mercer, E.H. and M.S.C. Birbeck. 1966. Electron
microscopy: A handbook for biologists. Blackwell
Scientific Publications. Oxford, England.
48.
McQuillen, Kenneth. 1960. Bacterial protoplasts.
p. 249-359. In I.C. Gunsalus and Roger Y. Stanier,
The bacteria. Volume I. Academic Press. New York.
49.
Noller, Eric C. and S.E. Hartsell. 1961. Bacteriolysis
of Entevobacteriaceae : Lysis by four lytic systems
utilizing lysozyme. J. Bacterid. 81:482-491.
50.
Panos, Charles. 1968. Comparative biochemistry of
membranes from Streptococcus pyogenes and derived
stable L-forms. p. 154-158. In Lucien B. Guze
(ed.), Microbial protoplasts, spheroplasts, and
L-forms. The Williams and Wilkins Co. Baltimore,
Maryland.
51.
Sagara, Yosunabu, Komei Fukui, Fusaoota, Nagayuki
Yoshida, Tatsuo Koshiyana, and Michimosa Fujimoto.
1971. Rapid formation of protoplasts of Streptomyces griseoflavus and their fine structure. Jap.
J. Microbiol. 15:73-84 .
52.
Salton, Milton R.J. 1968. Isolation and characterization of bacterial membranes, p. 144-153. In
Lucien B. Guze (ed.), Microbial protoplasts, spheroplasts, and L-forms. The Williams and Wilkins Co.
Baltimore, Maryland.
53.
Salton, Milton R.J. 1964. Isolation of bacterial cell
wall. p. 42-62. In The bacterial cell wall.
Elsevier Publishing Co. New York.
54.
Salton, M.R.J, and A.F.M. Ehtisham-ud-Din. 1965. The
localization of cytochromes and carotenoids in
isolated bacterial membranes and envelopes. Aust.
J. Exp. Biol. Med. Sci. £3:255-264.
55.
Salton, M.R.J, and J.H. Freer. 1965. Composition of
the membranes isolated from several gram-positive
bacteria. Biochim. Biophy. Acta. 107 :531-538 .
56.
Saperstein, S. and M.P. Starr. 1955. Association of
carotenoid pigments with protein components of
non-photosynthetic bacteria. Biochem. Biophy.
Acta. 16:482-488.
32
57.
Schnaitman, Carl. A. 1971. Effect of ethylenediaminetetraacetic acid, Triton X-100 and lysozyme
on the morphology and chemical composition of
isolated cell walls of Escherichia coli. J.
Bacteriol. 108:553-565.
58.
Schnaitman, Carl A. 1971. Solubilization of the
cytoplasmic membrane of Escherichia coli by
Triton X-100. J. Bacteriol. 108:545-552.
59.
Segen, Barbara J. and Kenneth D. Gibson. 1971.
Deficiencies of chromatophore proteins in some
mutants of Rhodopseudomonas spheroides with
altered carotenoids. J. Bacteriol. 105:701709 .
60.
Smith, P.F. 1963. The carotenoid pigments of Mycoplasma.
J. Gen. Microbiol. 32:307-319.
61.
Steensland, H. and H. Larsen. 1969. A study of the cell
envelop of the Balobacteria. J. Gen. Microbiol.
55^: 325-336 .
62.
Strominger, Jack L. 1968. Enzymatic reactions in
bacterial cell wall synthesis sensitive to penicillin and other antibacterial substances, p. 5561. In Lucien B. Guze (ed.), Microbial protoplasts,
spheroplasts and L-forms. The Williams and Wilkins
Co. Baltimore, Maryland.
63.
Szaniszlo, Paul J. 1968. The nature of intramycelial
pigmentation of some Actinoplanaceae. Jour. Elisha
Mitchell Sci. Soc. 1968:24-26.
64.
Takeya, Kenji and Kazuhito Hisatune. 1963. Mycobacterial
cell walls: I. Methods of preparation and treatment
with various chemicals. J. Bacteriol. 85:16-23.
65.
Taylor, Richard F., Ikawa Mujoshi, and William Chesbro.
1971. Carotenoids in yellow pigmented Enterococci.
J. Bacteriol. 105:676-678.
66.
Thirkell, D. and M.I.S. Hunter. 1969. Carotenoidglycoprotein of Sarcina flava membrane. J. Gen.
Microbiol. 58:289-292.
67.
Thirkell, D. and M.I.S. Hunter. 1970. A water soluble
carotenoid-glycopeptide from Sarcina morrhuae.
J. Gen. Microbiol. 62:125-127.
33
68.
Tsukamura, M. 1971. Proposal of a new genus Gordona
for slightly acid fast organisms occurring in sputa
of patients with pulmonary disease and in soil. J.
Gen. Microbiol. 68:15-26.
69.
Ungers, Grace E. and J.J. Cooney. 1968. Isolation and
characterization of carotenoid pigments of Micrococcus roseus. J. Bacteriol. 96 : 234 - 241.
70.
Wachsman, J.T. and R. Storck. 1960. Propionate
induced lysis of protoplasts of Bacillus megaterium.
J. Bacteriol. 80:600-606 .
71.
Warren, G.H., Jane Gray, and J.A. Yurchenco. 1957.
Effects of polymyxin on the lysis of Neisseria
catarrhalis by lysozyme. J. Bacteriol. 74:788789.
~~*
72.
Wayne, Lawrence G. 1967. Selection of characters for
an Adansonian analysis of Mycobacterial taxonomy.
J. Bacteriol. 93:1382-1391.
73.
Wayne, Lawrence G. and Wendy M. Gross. 1968. Base
composition of deoxyribonucleic acid isolated from
Mycobacteria. J. Bacteriol. 96 :1915-1919.
74.
Weedon, B.C.L. 1971. p. 29-60. Occurrence. In
Otto Isler (ed.), Carotenoids. Birkhauser~Verlag.
Stuttgart.
75.
White, David A., W.J. Lennarz, and Carl A. Schnaitman.
1972. Distribution of lipids in the wall and
cytoplasmic membrane subfractions of the cell
envelope of Escherichia coli . J. Bacteriol.
109:686-690.
76.
Work, Elizabeth and Hillary Griffiths. 1968. Morphology
and chemistry of cell walls of Micrococcus radiodurans. J. Bacteriol. 95:641-657.