Mycobacteria

zhanglu151

Function of Phosphatidylinositol in Mycobacteria
Phosphatidylinositol (PI) is an abundant phospholipid
in the cytoplasmic membrane of mycobacteria and
the precursor for more complex glycolipids, such as the
PI mannosides (PIMs) and lipoarabinomannan (LAM).
To investigate whether the large steady-state pools of PI
and apolar PIMs are required for mycobacterial growth,
we have generated a Mycobacterium smegmatis inositol
auxotroph by disruption of the ino1 gene. The ino1 mutant
displayed wild-type growth rates and steady-state
levels of PI, PIM, and LAM when grown in the presence
of 1 mM inositol. The non-dividing ino1 mutant was
highly resistant to inositol starvation, reflecting the
slow turnover of inositol lipids in this stage. In contrast,
dilution of growing or stationary-phase ino1 mutant in
inositol-free medium resulted in the rapid depletion of
PI and apolar PIMs. Whereas depletion of these lipids
was not associated with loss of viability, subsequent
depletion of polar PIMs coincided with loss of major cell
wall components and cell viability. Metabolic labeling
experiments confirmed that the large pools of PI and
apolar PIMs were used to sustain polar PIM and LAM
biosynthesis during inositol limitation. They also
showed that under non-limiting conditions, PI is catabolized
via lyso-PI. These data suggest that large pools of
PI and apolar PIMs are not essential for membrane integrity
but are required to sustain polar PIM biosynthesis,
which is essential for mycobacterial growth.
Mycobacterium tuberculosis, the causative agent of tuberculosis,
infects nearly one third of the world population and
causes active disease in an estimated 16 million people worldwide
(1). The distinctive cell wall of M. tuberculosis and other
pathogenic mycobacteria (M. leprae, M. avium-M. intracellulare
complex, and M. ulcerans) confers protection against a
range of microbicidal processes and many classes of antibiotics
and undoubtedly contributes to the success of these organisms
as pathogens. The mycobacterial cell wall contains a number of
unusual features, including the presence of a highly structured
peptidoglycan-arabinogalactan (AG)1-mycolic acid macromolecule
and a diverse array of glycolipids that form an asymmetric
outer bilayer with the mycolic acids (2–4). Mycobacteria and
other members of the Actinomycetales also differ from other
eubacteria in synthesizing phosphatidylinositol (PI) and the
biosynthetically related lipoglycans, PI mannosides (PIMs),
lipomannan (LM), and lipoarabinomannan (LAM) (5–7).
Whereas there is accumulating evidence that the PIMs and
LM/LAM have potent immuno-modulatory activities that may
be important for the pathogenesis of M. tuberculosis (7–10), the
presence of structurally related PIMs and LAMs in saprophytic
mycobacterial species suggests that these lipoglycans have a
more fundamental role(s) in mycobacterial physiology. This
conclusion is supported by the finding that both phosphatidylinositol
synthase and PimA, the first mannosyltransferase in
the PIM/LM/LAM pathway (Fig. 1), are essential for growth
and viability of the saprophytic mycobacteria species M. smegmatis
(11, 12). It is possible that the large steady-state pools of
PI and apolar PIM species may be required for cell membrane
integrity and/or other membrane functions. In this regard, it is
notable that PI is a bilayer-forming phospholipids, whereas
cardiolipin and phosphatidylethanolamine, the other major
phospholipids in the mycobacterial membrane, can form nonbilayer
structures (13). Alternatively, or in addition, PI and
apolar PIMs could function as a large dynamic pool of precursors
for polar PIMs and LM/LAM biosynthesis.
In order to investigate the potential role(s) of PI and PIMs as
both precursor and end products, we have generated a M. smegmatis
inositol auxotroph by disrupting the ino1 gene encoding
inositol-3-phosphate synthase (IPS). IPS is a key enzyme in the
only known pathway for the de novo synthesis of inositol from
D-glucose-6-phosphate (Fig. 1). The rate of synthesis of PI and
downstream products in this mutant should therefore be dependent
on the supply of extracellular inositol. Stoker and co-workers
(14) have recently disrupted the orthologous gene in M. tuberculosis.
The M. tuberculosis mutant required very high levels of
exogenous inositol for growth and was severely attenuated in
macrophages and severe combined immunodeficient (SCID) mice
(14). Surprisingly, cellular levels of inositol lipids did not decrease
when the M. tuberculosis mutant was suspended in inositol-
free medium, although this may have been due to the use of
non-replicating cultures (14). In this study, we show that inositol
starvation has little effect on the cellular levels of inositol lipids
or the viability of the non-replicating M. smegmatis ino1 mutant.
In contrast, dilution of the ino1 mutant in inositol-free medium
resulted in the rapid depletion of PI and apolar PIMs and a
slower depletion of polar PIMs. Depletion of polar PIMs coincided
with loss of other cell wall components and cell viability. We have
also identified a novel lipase-mediated catabolic pathway that is
involved in regulating cellular levels of PI in both wild-type and
mutant M. smegmatis. These findings provide new insights into
the function and metabolism of PI and PI-containing glycolipids
in mycobacteria.
EXPERIMENTAL PROCEDURES
Bacterial Strains and Growth Conditions—Escherichia coli XL-1
Blue MRF
(Stratagene) was grown in Luria Bertani (LB) medium.
Wild-type M. smegmatis mc2155 (15) and the mutants derived from it
were routinely grown on M9 agar (16) with 0.4% glucose and 1.5% agar
supplemented with 1 mM inositol, except where stated. Mycobacteria
used for all growth and biochemical studies were grown in M9 broth
with 0.4% (w/v) glucose and 0.05% (v/v) Tween 80 and supplemented
with inositol as required. Antibiotics, streptomycin (20
g/ml), kanamycin
(20
g/ml), or ampicillin (100
g/ml) was added to media as
required. Growth and survival of the cultures were measured by counting
colony-forming units (cfu) using solid M9 media containing 1 mM
inositol as a test medium or by measurement of cellular protein (17).
Genomic DNA was isolated from mycobacteria as described (18).
PCRs were performed in an MJ Research PTC-200 Peltier thermal
cycler. Reactions contained 10 ng of template DNA, 20 pmol of each
oligonucleotide primer, 2 units of Pfx polymerase (Invitrogen) or 0.2–1
unit of Taq polymerase (Roche Applied Science), and the enzyme reaction
buffer supplied by the manufacturer. Southern hybridization using
digoxygenin-labeled probes (Roche Applied Science) was performed according
to the manufacturer’s instructions. Transformation of bacteria
was performed using a Bio-Rad Gene Pulser. Competent M. smegmatis
cells were prepared as described previously (19).
Creation and Genetic Characterization of a M. smegmatis ino1 Mutant—
A Lambda FixII (Stratagene) genomic library of M. smegmatis
was screened by plaque hybridization using an oligonucleotide homologous
to the M. tuberculosis ino1 gene (5
-GACTTCCTGAACATGCTGG).
The probe bound to a phage clone, HBJ24, which had previously
been shown to contain ponA (20). Further analysis showed that
ino1 was a 1089-bp ORF that was contained on a 3.6-kb PstI fragment
along with three other ORFs.
The 3.6-kb PstI fragment was subcloned into the PstI site of pBluescript
SK() and then linearized by digestion of a BalI site 634 bp into
the ino1 ORF. A kanamycin resistance cassette, aphA-3, was derived by
PCR, using M13 universal primers, from pUC18K (21) and then digested
with SmaI and cloned into the BalI site of pHBJ228 to disrupt
ino1. This construct was then linearized by digestion of the HindIII site
of the pBluescript SK() cloning site, treated with T4-DNA polymerase,
and then ligated with a streptomycin resistance gene that had been
excised from pHP45 (22) by SmaI digestion. The resulting plasmid,pHBJ244, was transformed into M. smegmatis for allelic exchange to
replace the chromosomal copy of ino1 with a disrupted version. Transformants
were selected on LB media containing 55 mM inositol and
kanamycin. Potential double crossover mutants (kanamycin-resistant,
streptomycin-sensitive) were screened for their ability to grow in the
absence of inositol on M9 minimal medium. The disruption of ino1 in an
inositol auxotroph was confirmed by Southern hybridization. The blots
were hybridized with a 411-bp digoxygenin-labeled DNA probe derived
by PCR from ino1 using primers (5
-TCGGAAGAGGCCGACAAG and
5
-TTGCGGTCGTCGAGCCAG). The hybridizing PstI and Bgl11 fragments
of the mutant were 800 bp larger than the corresponding bands
in the parent, showing that ino1 had increased in size due to disruption
by the 800-bp aphA-3.
The ino1 mutant was complemented with an intact copy of the ino1
gene on a mycobacterial shuttle vector, pHBJ334, which is a streptomycin-
resistant derivative of pMV261 (23). The complementation
plasmid, pHBJ378, was made by cloning a PCR-amplified copy of the
ino1 gene (primers 5
-CGGAATTCTCTGAGCACGCAGGAG and 5
-
CGGAATTCAGCCCTCGATGAAGG) into the EcoRI site of the vector
such that the gene would be transcribed by the GroEL promoter. Both
pHBJ378 and the vector pHBJ334 were introduced into M. smegmatis
mc2155 wild type and ino1 mutant strains by electrotransformation (19).
IPS Enzyme Assay—The IPS activity was assayed essentially as
described previously (24), except that the inositol and glucose in the
samples were detected by aluminum-backed Silica Gel 60 high performance
thin layer chromatography (HPTLC) sheets (Merck). The HPTLC
was developed three times in 1-propanol:acetone:water (9:6:5, 5:4:1,
and 9:6:5, v/v), and then the amount of inositol and glucose was measured
on a HPTLC plate reader (Berthold). Enzyme activity was expressed
as units/mg protein. One unit was defined as the amount of
enzyme that catalyzed the formation of 1 nmol of D-myo-inositol-3-
phosphate in 1 h.
Extraction and Analysis of M. smegmatis Cell Wall Components—
Mycobacterial cells were harvested from liquid cultures by centrifugation
and washed in phosphate-buffered saline. Lipids were extracted
twice in 20 volumes of CHCl3:CH3OH (2:1, v/v) and once in
CHCl3:CH3OH:H2O (1:2:0.8, v/v) for 2 h at room temperature. The
extracts were combined and dried under a stream of N2, and lipids
were separated from salts and polar metabolites by biphasic partitioning
in 1-butanol:water (2:1, v/v). LM/LAM was extracted from the
delipidated pellet with three rounds of refluxing in 50% ethanol for
2 h at 100 °C. The combined 50% ethanol extracts were dried under
N2, and LAM was purified by octyl-Sepharose (Amersham Biosciences)
chromatography (25).
Lipids (PI, PIMs, and GPLs) were analyzed by HPTLC and loading
normalized to cellular protein (17). PI and PIMs were resolved in
CHCl3:CH3OH:1 M NH4OAc:13 M NH3:H2O (180:140:9:9:23, v/v) (solvent
1), and GPLs were resolved in CHCl3:CH3OH (9:1, v/v) (solvent 2).
HPTLC plates were stained with orcinol/H2SO4 (carbohydrate), cupric
acetate (lipids), or molybdate stain (phospholipids) (26). Cupric acetatestained
bands were quantitated by densitometry with the Kodak Digital
ScienceTM Image system (Eastman Kodak Co.). PI-specific phospholipase
C digestion was performed as described previously (27).
The monosaccharide composition of purified LAM fractions and AG
in the residual pellet was determined after hydrolysis in 2 M trifluoroacetic
acid (2 h, 100 °C) followed by conversion of the released
monosaccharides to their alditol acetate derivatives by reduction in
NaBD4 and acetylation in acetic anhydride using 1-methyl-imidazole
as a catalyst (28). Gas chromatography-mass spectrometry was performed
using a Hewlett Packard 6890 GC fitted with a HP-1 column
attached to a Hewlett Packard 5973 mass selective detector. For
analysis of alditol acetates, the column was held at 80 °C for 1 min,
ramped at 30 °C min1 to 140 °C and then at 5 °C min1 to 250 °C,
and held at 250 °C for 20 min.
Metabolic Labeling—Cells were grown to mid-exponential phase in
M9 broth with 1 mM inositol. Cells were washed twice in M9 broth
(37 °C) to remove the inositol. The concentrated cells (1:500 of original
volume) were pulsed with myo-[2-3H]inositol (50
Ci/ml) for 5 min at
37 °C. The cells were washed twice again in M9 broth (37 °C) to remove
the unincorporated myo-[2-3H]inositol before being resuspended in
fresh M9 media (37 °C) with 0 or 1 mM inositol at the same density as
when they were harvested. Growth of cultures was monitored by biomass
accumulation. Samples were harvested from the cultures during
the actively growing periods for analysis of cell wall components. Radiolabeled
lipids were detected by fluorography after coating the
HPTLC sheets with EA Wax (EA Biotech) and exposure to BioMax MR
film (Kodak) at 80 °C or scraped from the HPTLC sheets and label
quantitated by liquid scintillation counting.RESULTS
Characterization of the M. smegmatis ino1 Mutant—A gene
with 87% amino acid identity to the M. tuberculosis ino1 gene
was isolated from M. smegmatis. The M. smegmatis homologue
was present in an essentially identical loci to the M. tuberculosis
ino1 gene (29). Specifically, two ORFs of unknown function
(Rv0048c and Rv0047c in M. tuberculosis) were located
downstream of ino1, whereas another ORF of unknown function,
Rv0045c, precedes it. These ORFs have 48–81% amino
acid identity to M. smegmatis homologues. The protein encoded
by M. smegmatis ino1 had 87% amino acid identity to M. tuberculosis
IPS.
A M. smegmatis ino1 mutant was generated by targeted
disruption of the ino1 gene encoding a putative IPS (Fig. 2A).
Colonies of the ino1 mutant were only recovered from M9 agar
containing inositol, supporting a role for this gene in inositol
metabolism. Disruption of ino1 was confirmed by Southern
hybridization (Fig. 2B) and measurement of IPS activity.
Whereas wild-type M. smegmatis contained high levels of IPS
activity (3.29  0.77 units/mg protein), no IPS activity was
detected in the ino1 mutant (0.01  0.03 unit/mg protein).
Finally, complementation of the ino1 mutant with an intact
copy of the M. smegmatis ino1 gene on a mycobacterial shuttle
vector increased IPS activity to 0.15  0.08 unit/mg protein and
restored inositol prototrophy. These data indicate that ino1
encodes the only functional IPS in M. smegmatis and that the
ino1 mutant is auxotrophic for inositol.
Effect of Inositol Starvation on the Non-replicating ino1 Mutant—
Non-replicating stages of the ino1 mutant were highly
resistant to inositol starvation. When stationary-phase cells were
resuspended in inositol-free medium at high density, culture
viability remained constant over 400 h (Fig. 3A). Analysis of the
total lipid fraction showed that levels of polar PIMs (AcPIM6/
Ac2PIM6) did not decrease over this period, whereas levels of
apolar PIMs (AcPIM2/Ac2PIM2) decreased by 60% (Fig. 3B).
Remarkably, PI was not detectable in stationary-phase ino1 mutant,
regardless of whether inositol was present in the medium or
not (Fig. 3B). In contrast, wild-type M. smegmatis expresses high
levels of PI during stationary growth (Fig. 3B). These data suggest
that apolar PIMs are very stable metabolites that are turned
over very slowly in non-dividing cells. They also show that in the
absence of the endogenous pathway of inositol biosynthesis, stationary-
phase M. smegmatis is unable to accumulate PI. A large
steady-state pool of PI is thus not essential for the viability of
non-dividing M. smegmatis.
Effect of Inositol Starvation on Replicating Stages of the ino1
Mutant—To examine whether PI or downstream products were
essential for growth of M. smegmatis, stationary-phase ino1
cells were diluted into fresh medium containing 1000
M inositol,1
M inositol, or no inositol. The growth kinetics of the
ino1 mutant was indistinguishable from that of wild-type
M. smegmatis in the presence of 1000
M inositol (Fig. 4, A and
B; data not shown). In contrast, biomass did not increase substantially
(0.5 log unit), and viability decreased to zero over
200 h when the ino1 mutant was diluted into inositol-free
medium (Fig. 4, A and B). Interestingly, ino1 mutant cells
remained intact for 25 days after loss of viability, as shown by
unchanged biomass (Fig. 4A). The ino1 mutant grew slowly in
the presence of 1
M inositol, and the viability of these cultures
decreased dramatically after 150 h (Fig. 4, A and B).
Aliquots of the ino1 mutant grown in the presence of 1 or
1000
M inositol were harvested at various time points to
determine whether the loss of viability of cultures grown in 1

M inositol was associated with the loss of specific inositol
lipids. PI was detected in inositol-replete cultures during exponential
growth phase but decreased to below the level of detection
in stationary-phase cultures (Fig. 5A). In contrast, PI
never accumulated to detectable levels in either exponential or
stationary-phase ino1 cells grown in the presence of 1
M
inositol (Fig. 5A). These data extend the observation made on
stationary-phase cells, suggesting that a large pool of PI is not
required for either the viability or growth of M. smegmatis.
Because wild-type mycobacteria express high levels of PI
throughout growth (Fig. 3B), these data also suggest that theino1 mutant has a reduced capacity to take up inositol in
stationary phase and/or an increased dependence on endogenously
synthesized inositol for PI biosynthesis.
Unlike PI, cellular levels of PIM2 and PIM6 were expressed
throughout growth when the ino1 mutant was grown in 1000

M inositol, with the relative abundance of the diacylated species
increasing with culture age (Fig. 5B). However, in the
presence of 1
M inositol, levels of apolar PIM species decreased
to below the level of detection within 78 h. Levels of polar PIMs
also decreased, but at a slower rate (Fig. 5B). Surprisingly,
LAM levels in inositol-deprived ino1 mutant decreased to
20% of the level found in the ino1 mutant grown in 1000
M
inositol, but then they remained constant for the remainder of
the experiment (Fig. 5C). The initial decrease in LAM levels
may reflect the dilution of this component as a result of ongoing
cell division (Fig. 4). The fact that apolar and polar PIMs
persisted for longer than LAM suggested that most of the PI
was redistributed into PIM biosynthesis rather than LAM biosynthesis
under inositol-limiting conditions.
To investigate whether the loss of polar PIMs was associated
with loss of other cell wall components, the ino1 mutant was
grown in 1000 or 1
M inositol, and cells were harvested at the
beginning of stationary phase. As shown in Fig. 5, D and E,
components of both the outer lipid layer (GPLs) and inner
peptidoglycan-AG-mycolic acid complex (AG) were reduced by
90% in inositol-limited cultures. In contrast, cellular levels of
cytoplasmic membrane phospholipids, phosphatidylethanolamine,
and cardiolipin were not decreased after inositol starvation
(Fig. 5A). These data suggest that the loss of polar PIMs
has little effect on the composition of the cytoplasmic membrane
but is associated with the catastrophic loss of major cell
wall components and cell viability.
Relationship between PI and PIM in Actively Dividing Mycobacteria—
The affect of inositol starvation on actively dividing
mycobacteria was monitored by cultivating the ino1 mutant
in 1000
M inositol and then transferring cells in mid-exponential
phase to fresh medium containing either 1000
M inositol
or no inositol at the same cell density. M. smegmatis wild typeand the ino1 mutant in 1000
M inositol medium continued to
multiply for 15 h before entering stationary phase (Fig. 6A).
Whereas levels of PI decreased by 50% when the ino1 mutantreached stationary phase, cellular levels of other phospholipids
and total PIM remained constant throughout exponential and
stationary growth (Fig. 7, A and C). Interestingly, levels of
diacylated PIM2 and PIM6 in the ino1 mutant (Fig. 7C) and
wild type (data not shown) invariably increased at 5 h and
subsequently decreased, suggesting that the inositol acylation
reaction is reversible. In the absence of exogenous inositol, the
actively dividing ino1 mutant underwent 3–4 cell divisions
before biomass plateaued after 15 h and viability decreased
after 26 h (Fig. 6B). Inositol starvation led to the rapid decrease
in PI (depleted within 3 h) and apolar PIMs (depleted within 5
h) (Fig. 7, B and D) but to a small increase in AcPIM6 levels
over 5 h (Fig. 7, B and D). Levels of LM/LAM remained constant
during this period (data not shown). These data support
the notion that the major cellular pools of PI and PIM2 can be
used to sustain polar PIM synthesis during nutrient starvation.
They also suggest that dividing cells are more resistant to
inositol starvation than stationary-phase cells, by virtue of
containing a larger steady-state pool of PI.
Catabolism of PI during Inositol Starvation—The analyses
described above suggest that the large cellular pools of PI and
PIM2 are dynamic and rapidly channeled into polar PIM/LM/
LAM and/or are catabolized during inositol starvation. To investigate
which of these processes predominates under inositolreplete
or -limiting conditions, M. smegmatis wild type and the
ino1 mutant were pulse-labeled with myo-[3H]inositol and then
resuspended in fresh medium containing 1000
M inositol or no
inositol. In wild-type M. smegmatis, label was initially incorporated
into PI and subsequently chased into AcPIM2, LAM,
and AcPIM6 (Fig. 8, AD). The efficiency of labeling of the
latter species was increased when the chase was performed ininositol-free medium, which appeared to coincide with reduced
turnover of labeled PI (Fig. 8, A and C). Regardless of the chase
conditions, total incorporation of myo-[3H]inositol into PIM and
LAM fractions never exceeded 48–56% of the label initially
incorporated into the PI fraction (Fig. 8, B and D). These data
suggest that the majority of the PI may be hydrolyzed rather
than incorporated into PIMs and LAM, and the rate of hydrolysis
appears to be reduced when exogenous inositol levels are
low. To investigate whether PI turnover and the channeling of
PI into PIM biosynthesis is regulated by inositol availability,
identical labeled experiments were performed with the ino1
mutant. As expected, inositol phospholipids were labeled more
efficiently in the ino1 mutant because exogenous label was not
being diluted by de novo synthesized inositol (Fig. 8, E and F).
As observed in wild-type cells, labeled PI was rapidly turned
over when the ino1 mutant was suspended in high-inositol
medium (Fig. 8, E and F). Approximately 42% of the label in the
PI fraction was initially chased into AcPIM2 and subsequently
chased into AcPIM6. A novel species with a slower HPTLC
mobility than PI or AcPIM2 was also observed at the end of the
pulse and throughout the chase (Fig. 8E). This species was
susceptible to PI-specific phospholipase C digestion and mild
base hydrolysis and comigrated with authentic lyso-PI (data
not shown). This species was also detected in wild-type cells,
when fluorographs were developed for longer periods (data not
shown). When the ino1 mutant pulse-labeled with [3H]inositol
was chased in inositol-free medium, there was a marked increase
in the extent to which labeled PI was chased into
AcPIM2 and AcPIM6 (73%) and a reduction in the rate at
which label was chased out of the lyso-PI species (Fig. 8, G and
H). These analyses demonstrate that the entire pool of newly
synthesized PI in both the wild type and ino1 mutant can be
utilized for PIM and LM/LAM synthesis or catabolized via
lyso-PI. The catabolism of PI is dramatically reduced when
inositol is limiting, presumably reflecting the channeling of
limiting PI precursors into PIM and LM/LAM synthesis.DISCUSSION
We have generated a M. smegmatis inositol auxotroph by
disrupting the ino1 gene and examined the consequences of
inositol starvation on the synthesis and turnover of inositol
lipids and cell viability. Our analyses show that the PI pool of
M. smegmatis is very dynamic and rapidly depleted under
inositol-limiting conditions. Cells remain viable without PI,
although PI depletion eventually leads to depletion of polarPIMs, loss of other cell wall components, and cell death. A
major function of the large cellular pool of PI is thus to provide
a dynamic pool of precursors for polar PIM and LM/LAM biosynthesis.
We also show that the steady-state levels of PI may
be regulated by the action of one or more PI-specific lipases.
The growth phenotype of the M. smegmatis ino1 mutant is
similar to that of the recently generated M. tuberculosis ino1
mutant, which was unable to survive in macrophages or highly
susceptible SCID mice (14). Both mutants required exogenous
inositol for growth but survived for extended periods of time
without inositol while in stationary growth phase (this study
and Ref. 14). The resistance of stationary-phase cultures to
inositol starvation appears to reflect the lower requirement for
new membrane and cell wall components in non-dividing
stages and is consistent with the finding that the rate of PIM
synthesis is dramatically down-regulated in stationary-phase
M. smegmatis.2 Interestingly, the growth of the M. smegmatis
ino1 mutant was the same as that of wild-type cells in the
presence of 1 mM inositol, whereas the M. tuberculosis ino1
mutant required much higher levels of inositol (70 mM) for
normal growth (14). M. tuberculosis may be less efficient at
scavenging inositol from the environment than the saprophytic
M. smegmatis and/or have a higher dependence on inositol
generated via the D-glucose-6-phosphate pathway.
A dramatically different response was observed when either
stationary-phase or mid-exponential phase ino1 mutant cells
were diluted into inositol-free medium. Under these conditions,
the steady-state levels of PI and apolar PIMs decreased rapidly.
Metabolic labeling experiments indicated that the decrease
in PI and apolar PIMs was mainly due to the conversion
of these lipids into polar PIMs. Significantly, PI levels decreased
to below the level of detection, while inositol-starved
cells were still in mid-exponential growth. Taken together with
the observation that stationary-phase ino1 mutant cells lacking
detectable PI remain viable for extended periods, these
data suggest that large steady-state pools of PI and apolar
PIMs are not essential for either the viability or growth of
M. smegmatis. In contrast, the cellular levels of polar PIM
species (i.e. AcPIM6 and Ac2PIM6) decreased slowly during
inositol limitation, and loss of these glycolipids was associated
with the catastrophic loss of other cell wall components (AG,
outer layer glycolipids) and cell viability. Interestingly, cellular
levels of LM/LAM decreased more rapidly than polar PIMs
during initial stages of inositol limitation but then stabilized at
20% of wild-type levels when cells stopped dividing. The
decrease in LM/LAM may thus represent the dilution of these
molecules during cell division. The slower decrease in polar
PIM levels during inositol limitation also suggests that apolar
PIMs are preferentially directed into apolar PIM synthesis
rather than LAM synthesis during inositol starvation. Collectively,
these data suggest that the large steady-state pools of PI
and apolar PIMs are primarily metabolic precursors that can
be used to sustain polar PIM synthesis when mycobacteria are
subjected to nutrient limitation. It remains unclear whether
the loss of viability of inositol-starved cells is due to loss of
specific polar PIM species or the global loss of all inositol lipids.
M. smegmatis and M. marinum mutants with defects in apolar
PIM biosynthesis have recently been generated (30).3 These
mutants display a relatively mild growth and colony phenotype
but also accumulate high levels of PIM precursors (30). Thus,
whereas polar PIMs may be important for cell wall biogenesisand cell viability, these functions may also be fulfilled by
smaller PIM species.
The phospholipids, PI, phosphatidylethanolamine, and cardiolipin,
are thought to be restricted to the cytoplasmic membrane
(2). In contrast, it has been proposed that the PIMs may
be located in the cytoplasmic membrane and/or transported to
the cell wall or extracellular space (31, 32). The finding that the
entire pool of Ac/Ac2PIM2 can be utilized as precursors for
polar PIM synthesis suggests that these glycolipids are not
transported to the outer layer of the cell wall but remain within
the cytoplasmic membrane and/or cell fractions that contain
enzymes involved in polar PIM biosynthesis. At least one of
these enzymes, PimC, a putative mannosyltransferase that
catalyzes the conversion of AcPIM2 to AcPIM3, appears to be
located in the cytoplasmic membrane, based on the finding that
it utilizes the cytoplasmic mannose donor, GDP-mannose, and
lacks a recognizable signal sequence (33). We have also obtained
evidence that the polyprenol-phosphate-mannosedependent
mannosyl-transferases involved in the synthesis of
polar PIMs are also located in the cytoplasmic membrane (25).
Collectively, these data suggest that the major pools of apolar
PIMs remain in the cytoplasmic membrane.
The finding that PI was rapidly and constitutively catabolized
in M. smegmatis was unexpected. PI catabolism was observed
in both M. smegmatis wild type and the ino1 mutant
and appeared to involve the generation of a lyso-PI species.
Mycobacteria express a number of phospholipase activities,
although the endogenous or exogenous targets of these activities
are not well defined (34–36). The identification of lyso-PI
species implies the involvement of a PI-specific PLA2, although
phospholipase C or D activities could also be involved and
would not be detected in these analyses. Interestingly, PI catabolism
was reduced when cells were suspended in inositolfree
media, suggesting that the activity of the lipases can be
regulated by inositol availability and/or that the PimA mannosyltransferase
is more efficient at utilizing limiting pools of PI.
Collectively, these data indicate that the primary role of PI in
mycobacteria is to act as a dynamic pool of precursors for PIM
and LM/LAM synthesis and that the steady-state level of this
phospholipid is regulated by the rate of biosynthesis, the conversion
of PI to PIMs, and lipase-mediated catabolic reactions.