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J Biol Chem, Vol. 274, Issue 51, 36579-36584, December 17, 1999
From the Departments of Deacylation of purified
lipopolysaccharides (LPS) markedly reduces its toxicity toward mammals.
However, the biological significance of LPS deacylation during
infection of the mammalian host is uncertain, particularly because the
ability of acyloxyacyl hydrolase, the leukocyte enzyme that deacylates
purified LPS, to attack LPS residing in the bacterial cell envelope has
not been established. We recently showed that the cellular and
extracellular components of a rabbit sterile inflammatory exudate are
capable of extensive and selective removal of secondary acyl chains
from purified LPS. We now report that LPS as a constituent of the
bacterial envelope is also subject to deacylation in the same
inflammatory setting. Using Escherichia coli LCD25, a
strain that exclusively incorporates radiolabeled acetate into fatty
acids, we quantitated LPS deacylation as the loss of radiolabeled
secondary (laurate and myristate) and primary fatty acids
(3-hydroxymyristate) from the LPS backbone. Isolated mononuclear cells
and neutrophils removed 50% and 20-30%, respectively, of the
secondary acyl chains of the LPS of ingested whole bacteria. When
bacteria were killed extracellularly during incubation with ascitic
fluid, no LPS deacylation occurred. In this setting, the addition of
neutrophils had no effect, but addition of mononuclear cells resulted
in removal of >40% of the secondary acyl chains by 20 h.
Deacylation of LPS was always restricted to the secondary acyl chains.
Thus, in an inflammatory exudate, primarily in mononuclear phagocytes,
the LPS in whole bacteria undergoes substantial and selective
acyloxyacyl hydrolase-like deacylation, both after phagocytosis of
intact bacteria and after uptake of LPS shed from extracellularly killed bacteria. This study demonstrates for the first time that the
destruction of Gram-negative bacteria by a mammalian host is not
restricted to degradation of phospholipids, protein, and RNA, but also
includes extensive deacylation of the envelope LPS.
The prominent role ascribed to
LPS1 in evoking beneficial as
well as harmful mammalian host responses to invading Gram-negative bacteria has prompted intense scrutiny of host recognition,
detoxification, and elimination of this bacterial product.
Underacylated and underphosphorylated biosynthetic precursors and
chemically synthesized analogs of lipid A (the endotoxic part of LPS)
possess greatly diminished endotoxic activity and can act as
antagonists in human cell bioassays (1, 2). Mammalian enzyme systems
capable of removing acyl and phosphate groups from lipid A may
therefore play an important role in host defense against Gram-negative
bacteria and their LPS. Thus far, only one mammalian enzyme,
acyloxyacyl hydrolase, has been shown to deacylate LPS (3, 4).
We recently showed that both the cells and extracellular fluid of a
sterile inflammatory exudate actively deacylate purified LPS ex
vivo in an acyloxyacyl hydrolase-like fashion (5). It has been
much more difficult to analyze LPS breakdown products when the LPS is
presented in intact bacteria, because methods for selectively labeling
the acyl chains of LPS during their biosynthesis do not exist and
because the fatty acids released from LPS by leukocytes are rapidly
incorporated into cellular lipids (6). As a result, it is still unknown
whether the mammalian host is also capable of degrading and detoxifying
LPS in its natural setting, the bacterial outer membrane. This is
especially important because whole bacteria can signal mammalian host
cells with greater potency than isolated LPS (7) and because
introduction of purified LPS and live bacteria into animals yield
different cellular distributions and subsequent fates of
immunologically detectable LPS (8).
We now describe methods that allow quantitative separation of all of
the major classes of radiolabeled bacterial molecules and their
degradation products, permitting for the first time a reliable
quantitative assessment of LPS deacylation during extracellular and
intracellular antibacterial action of the mammalian host. We show that
in this inflammatory setting, loss of bacterial viability was not only
accompanied by hydrolysis of large portions of the bacterial
phospholipids, RNA, and protein (6, 9, 10) but also by extensive
deacylation of LPS by an acyloxyacyl hydrolase-like activity, chiefly
in the mononuclear phagocytes in the exudate.
Materials
Sodium [2-14C]acetate (57 mCi/mmol),
L-[35S]methionine (1175 Ci/mmol),
[2-14C]uracil (2.25 mCi/mmol) were from NEN Life Science
Products. Human serum albumin (HSA) was from Armour Pharmaceutical Co.
(Kanakee, IL). Oyster glycogen was from U. S. Biochemical Corp.
Hanks' balanced salt solutions without phenol red, with
(HBSS+) and without (HBSS Bacterial Strains and Growth Conditions
E. coli LCD25, an aceEF, gltA
strain of E. coli K12 unable to produce acetate or use
acetate as a carbon or energy source, was grown in modified minimal E
medium (12) with twice the stated concentrations of salts, glucose, and
amino acids as described by Munford et al. (11) with the
following modifications. Bacteria were grown overnight at 30 °C in
liquid 2× E medium containing 0.8 mM sodium acetate,
sedimented in a microcentrifuge at 3000 × g, washed
once in sterile physiological saline, and then subcultured for 20 h at 30 °C at a starting concentration of 2.4 × 107 bacteria/ml. Bacterial LPS and PL were labeled using
0.3 mM (17 µCi/ml) sodium [2-14C]acetate,
yielding ~15,000 cpm incorporated/106 bacteria. Protein
and RNA were labeled using the growth conditions described above,
except subcultures were grown with 0.3 mM unlabeled sodium
acetate and were supplemented with 5.5 µCi/ml
L-[35S]methionine (yielding 700 to 2000 cpm/106 bacteria) or 2 µCi/ml 2-[14C]uracil
(yielding 300 cpm/106 bacteria). After labeling, the
bacteria were sedimented in a Microfuge at 3000 × g
and cultured an additional 30 min at 37 °C in an equal volume of
fresh modified E medium containing 0.8 mM unlabeled sodium
acetate to drive cell-associated radiolabeled precursors into
macromolecules. Sterile HSA was added to 1% (to remove unesterified
fatty acids), and the bacteria were harvested by centrifugation at
3000 × g.
Collection and Separation of Rabbit Leukocytes and Ascitic Fluid
(AF)--
Sterile inflammatory peritoneal exudates were collected from
New Zealand White rabbits, and cells (PMN and mononuclear cells (MNC))
and AF were isolated as described previously (5). To provide a
biological fluid containing opsonins but devoid of antibacterial activity, AF from complement (C6)-deficient rabbit exudates was incubated with CM-Sephadex resin to remove cationic proteins as described previously (13), yielding unbound C6-deficient AF (C6d UBAF).
Assay Conditions
Labeled E. coli LCD25 were incubated (final
concentration of 1 × 108/ml) with 90% (v:v) AF alone
or with rabbit exudate cells as indicated in 90% (v:v) AF or 50%
(v:v) C6d UBAF in HBSS+ containing 1% HSA. All incubations
(200-µl volume) were buffered to pH 7.3-7.4 with 20 mM
HEPES. For protein degradation experiments, incubations with leukocytes
contained 0.5 mM cycloheximide to inhibit potential host
cell reincorporation of labeled amino acids from degraded bacterial
protein (9). The leukocyte concentration was 1 × 107/ml for protein and RNA degradation and 1.5-2.0 × 107/ml for LPS and phospholipid degradation. Incubations
proceeded at 37 °C with gentle (100 rpm) shaking in air for 1 h
and thereafter at 37 °C without shaking in 5% CO2 to
maintain pH.
Protein and RNA Degradation
To ensure effective resuspension of clumped leukocytes,
supernatant fluid (200 µl) was removed from tubes and replaced with 300 µl of divalent cation-poor buffer (HBSS Lipid Analysis (see Fig. 1)
At the indicated times, samples containing bacteria labeled with
sodium [2-14C]acetate were extracted according to the
procedure of Bligh and Dyer (14). At time = 0, mean partitioning
(± S.E.) of the total bacterial radiolabeled material in this step was
74.6 ± 0.8% into chloroform, 23.9 ± 0.8% into the
interface (IF), and 1.5 ± 0.2% into H2O/methanol,
with essentially complete recovery of cpm.
A) Phospholipid Degradation--
After growth of the bacteria
with [14C]acetate, 68% of the total incorporated cpm
were in the PLs. Intact PLs and the radiolabeled products of PL
degradation were quantitatively recovered in the CHCl3
phase of the initial Bligh and Dyer extraction. Degradation of
bacterial PL was measured using a two-step TLC procedure. First, radiolabeled lipids in the CHCl3 phase were resolved by TLC
on silica gel G using solvent system 1 (TLC1, see below).
Intact bacterial PL remained at the origin in this system, along with other radiolabeled species including partially deacylated bacterial PL
(lyso-PG and lyso-PE) and leukocyte PL (mainly PC, radiolabeled by
incorporation of labeled FFAs released from bacterial lipids). These
lipids were eluted from the silica and resolved by TLC on silica gel G
using solvent system 2 (TLC2, see below). Radiolabeled species on all TLC plates were detected and quantitated by proportional argon ionization using an AMBIS-1000 detector (Ambis Inc., San Diego,
CA). Radiolabeled species were identified by comparison with the
migration of unlabeled authentic standards (NFA (18:1), PE, PG, CL,
LPG, LPE) detected by staining with iodine vapor.
In unincubated (time = 0) bacteria, B) LPS Deacylation--
Intact and partially deacylated LPS were
quantitatively recovered at the IF of the initial Bligh and Dyer
extraction (see Fig. 1, Table IA and Ref.
5). LPS deacylation was measured as the loss of radiolabeled
LPS-derived fatty acids (3-OH-14:0, 12:0, and 14:0) from the interface.
After release from the IF, these fatty acids accumulated in the
chloroform phase as FFAs or fatty acids esterified into host cell
lipids (5).
To measure the fatty acid composition of the LPS collected from the IF,
this material was dried under N2 and hydrolyzed with 4 N HCl and 4 N NaOH as described previously
(15). The samples were then acidified to pH 4.0 with acetic acid and
extracted by the method of Bligh and Dyer. Of the cpm recovered after
hydrolysis of the IF, 95.8 ± 1.2% was recovered after
re-extraction. Of these cpm, 92-100% partitioned into the chloroform
phase. The LPS-derived FA were then resolved and quantitated using TLC
in two steps. First, the hydroxylated (3-OH-14:0) and nonhydroxylated
(NFA) (12:0, 14:0) fatty acids were separated by TLC1.
Second, to resolve 12:0 and 14:0, the NFA from TLC1 were
eluted from the silica gel and separated by reverse-phase TLC using
KC18-substituted silica gel and solvent system 3 (TLC3, see
below). Labeled species on all TLC plates were detected and quantitated
by AMBIS-1000 or by autoradiography. Labeled FFA standards (3-OH-14:0
derived from purified 14C-acetate-labeled E. coli LPS after acid/base hydrolysis and [14C]12:0,
[3H]14:0, and [14C]16:0 obtained from
commercial sources) were run on TLC in parallel to identify labeled
species from experimental samples. The total cpm in 3-OH-14:0 in the
interface at a given time was calculated as follows: (total cpm in
hydrolyzed IF) × (% re-extracted into CHCl3) × (% of this material migrating as 3-OH-14:0 in TLC1).
The total cpm in 12:0 or 14:0 in the interface at a given time was
calculated as follows: (total cpm in hydrolyzed IF) × (% re-extracted into CHCl3) × (% of this material
migrating as NFA in TLC1) × (% of this material
migrating as either 12:0 or 14:0 in TLC3).
LPS deacylation is expressed as % loss of each fatty acid from the
interface over time, using the values for each acyl species obtained
from unincubated bacterial suspensions (time = 0) as 100%.
TLC--
Three TLC systems were used, as described previously
(16).
TLC1
TLC2
TLC3 Elution from Silica G--
After visualization with AMBIS-1000,
radioactive spots of silica G were scraped from the plate and eluted by
washing once with 200-500 µl of chloroform/methanol/acetic
acid/H2O (55:33:9:4; v:v:v:v), once with an equal volume of
chloroform, and once again with chloroform/methanol/acetic
acid/H2O (55:33:9:4; v:v:v:v). Each wash step was preceded
by a 10-min incubation at 42 °C. Recovery of radiolabeled lipids
after elution was 83.7 ± 1.5% and was essentially the same for
PL and various FA species.
Uptake of Bacteria by Host Cells--
After a 1-h incubation
(see assay conditions), samples containing bacteria and leukocytes were
centrifuged at 70-100 × g to separate
leukocyte-associated and extracellular bacteria. The leukocyte pellet
was washed once with HBSS+ containing 0.1% HSA and once
with HBSS Bacterial Viability--
After 1 h at 37 °C, samples
were taken, diluted with sterile saline, and plated on nutrient broth
agar as described previously (13). Viability was measured as the number
of colonies formed after incubation at 37 °C for 18-24 h.
Quantitative Recovery of LPS from E. coli LCD25 and Separation from
PL--
To measure deacylation of LPS in E. coli LCD25
during extracellular killing by AF and intracellular killing by
inflammatory exudate cells, it was necessary to resolve LPS-derived
fatty acids from FA linked to other bacterial macromolecules
(principally phospholipids) that were labeled during growth in
[14C]acetate-supplemented medium. Acetate-labeled LPS
purified from E. coli LCD25 contained mainly three fatty
acid species: 3-OH-14:0 (see Fig. 2A, lane 2),
12:0 and 14:0 (Fig. 2B, lane 3) (11); see
also Ref. 5. All 3-OH-14:0 and nearly all 12:0 in E. coli is
found in the lipid A moiety of LPS, whereas 14:0 in E. coli is divided roughly equally between PL and LPS (17).
Using the extraction procedure of Bligh and Dyer (see Fig.
1), essentially all compounds in E. coli LCD25 that contained labeled 3-OH-14:0 were recovered in the
IF between the aqueous and CHCl3 phases (Table IA and Fig.
2A, lanes 3 and
4), and virtually none were recovered in the
CHCl3 phase (Table IA and Fig. 2A, lane 5). Since 3-OH-14:0 is unique to the lipid A moiety of bacterial LPS, this finding indicates that nearly all bacterial LPS was recovered
in the IF. Similar complete recovery of partially deacylated LPS in the
IF was shown recently (5). In contrast, during this extraction
procedure, Determination of PL Hydrolysis--
For analysis of deacylation of
phospholipids, we used the chloroform phase of the same extracts of
cell suspensions that provided the IF for determination of LPS
deacylation. In TLC1, intact PL and lyso-PL remained at the
origin of the chromatogram (Fig. 2A, lanes 6-9).
The percentage of this radiolabeled material that consisted of intact
bacterial PL (PE, PG, or CL) was determined using TLC2. By
20 h in AF (Fig. 2A, lane 7), the labeled
species at the origin of TLC1 represented almost
exclusively monoacyl catabolites of bacterial PL (LPE and LPG (not
shown)), whereas after ingestion by PMN (Fig. 2A, lane
8) or MNC (Fig. 2A, lane 9), some of the
label at the origin was contained in the phagocyte lipid PC (not
shown). Labeled species that co-migrated with the solvent front also
accumulated; these were probably neutral lipids (e.g.
triglycerides), reflecting leukocyte incorporation of labeled FA that
were released from bacterial lipids, as reported previously (3,
6, 10) and in a recent publication (5).
Deacylation of LPS and PL during Extracellular Killing of E. coli
by AF--
To examine LPS deacylation during extracellular bacterial
killing, [14C]acetate-labeled E. coli LCD25
were incubated with 90% AF, resulting in rapid killing of E. coli LCD25 ( Leukocyte-dependent Deacylation of LPS in E. coli
Killed Extracellularly by AF--
We have shown recently that
acyloxyacyl hydrolase activity is much greater in inflammatory
exudate leukocytes than in AF (5). Therefore, LPS deacylation may
require either uptake of LPS shed from killed bacteria or ingestion of
intact bacteria. When bacteria were incubated with inflammatory exudate
leukocytes in the presence of AF, bacterial killing and PL hydrolysis
took place principally extracellularly, i.e. before
ingestion. This was evident from our inability to observe intact
bacteria inside phagocytes by light microscopy (not shown). Moreover,
the rate and extent of bacterial PL hydrolysis by AF plus phagocytes
paralleled much more closely the effects of AF alone than those of PMN
and MNC toward ingested bacteria (compare Figs. 3 and
4). The addition of MNC but not of PMN to
AF resulted in substantial loss of 12:0 and 14:0 from bacterial LPS
(Fig. 3), in a pattern closely similar to that seen when these
cells take up purified LPS (5).
LPS and PL Deacylation during Intracellular Killing by Inflammatory
Exudate Leukocytes--
Incubation of
[14C]acetate-labeled E. coli LCD25 with either
PMN or MNC in the presence of C6-deficient UBAF, which lacks
antibacterial activity (not shown) but contains complement-derived
opsonins present in the exudate, resulted in substantial bacterial
ingestion. After 1 h, 61% of the radiolabeled bacteria were
associated with PMN, which contained an average of 4 bacteria/cell (as
judged by light microscopy; not shown). Similarly, MNC took up 65% of bacterial radiolabel after 1 h, with an average of 7 bacteria/cell. Accompanying this uptake, PMN and MNC caused 99 and 86%
loss of bacterial colony-forming units within 1 h and up to 70 and
90% hydrolysis of bacterial PL, respectively, within 8-20 h (Fig. 4).
PMN caused significant (albeit slower and less extensive) loss of 12:0
and 14:0 from bacterial LPS reaching 20-30% by 20 h (Fig. 4).
MNC more rapidly and extensively deacylated LPS, as is shown by the
loss of nearly 50% of the 12:0 and 14:0 from the interface by 4 h. No deacylation occurred in C6-deficient UBAF alone (data not shown).
Thus, whereas only MNC effectively deacylated cell-free LPS after
extracellular killing of bacteria by inflammatory fluid, PMN as well as
MNC caused LPS deacylation after ingestion of intact bacteria. Under
all conditions tested, 3-OH-14:0 was quantitatively retained in the
interface (Figs. 3 and 4), demonstrating that the acyloxyacyl bonds of
lipid A were the sole target of the LPS-deacylating activity manifest
during and after killing of E. coli by inflammatory exudates.
Degradation of RNA and Proteins--
The preceding findings show a
dramatic difference in the deacylation of PL and LPS during killing by
either AF or PMN. To determine whether degradation of bacterial
macromolecules under these conditions is generally slow and limited (as
for LPS) or more rapid and extensive (as for PL), degradation of
biosynthetically labeled bacterial RNA and proteins was determined
during extracellular killing of E. coli LCD25 by AF or
during intracellular killing by inflammatory exudate leukocytes ( Previous analysis of the fate of LPS of whole bacteria has been
hampered until recently by technical difficulties including radiolabeling of LPS to adequate specific activity and separation of
LPS and its degradation products from other radiolabeled bacterial constituents. It has been claimed in previous studies that mouse macrophages can slowly degrade LPS of biosynthetically labeled bacteria
(20, 21). However, because reaction products could not be identified or
chemically characterized, the nature and extent of the process
remained undefined.
The introduction of E. coli LCD25 for high specific activity
fatty acid radiolabeling with acetate (11) has now made possible reliable detection of lipid A-derived degradation products (FA and
partially deacylated LPS) using as few as five bacteria per host cell.
Incorporation of acetate by this E. coli mutant is limited
to fatty acyl groups of LPS, phospholipids, and lipoproteins. Virtually
complete separation of these bacterial products and their constituent
fatty acids can be accomplished by the methods used in this study. Our
ability to assess the recovery of LPS and the radiological purity of
the recovered LPS is based upon the labeling of 3-OH-myristate, a
unique and quantitatively constant acyl constituent of lipid A.
Deacylation of LPS was determined by measuring the loss of the lipid
A-specific fatty acids 3-OH-14:0 and 12:0 from the LPS-containing IF
between the aqueous and chloroform phases of extracted samples. Because
the IF is essentially free of phospholipids (18), deacylation of lipid
A was also assessed by measuring the loss of 14:0, a major fatty acyl
constituent of lipid A and only a minor component of phospholipids. In
the IF of extracts of bacteria, the appearance of material containing
radiolabeled species co-migrating with 16:0 (Fig. 2B) may be
attributed to the presence of Braun lipoproteins containing labeled
acyl chains. The acyl groups in this lipoprotein (75% 16:0 and 18:1)
(22) co-migrate with 16:0 and not with any of the fatty acyl groups of
lipid A, and therefore do not confound the analysis of LPS deacylation.
During incubation with ascitic fluid, prompt killing of E. coli LCD25 was observed (not shown) in accord with the known
antibacterial properties of rabbit inflammatory peritoneal fluid (13).
Killing was accompanied by extensive disassembly of the bacteria
indicated by complete hydrolysis of PL (Fig. 3), extensive degradation
of RNA (Fig. 5), and lack of uptake of visibly intact bacteria by phagocytes (not shown). However, no LPS deacylation was observed (Fig.
3A), despite the ability of ascitic fluid to degrade
purified LPS (5). LPS still associated with bacterial remnants or with bacterial outer membrane proteins or phospholipids (23-25) may be less
subject to deacylation than purified LPS. Alternatively, because the
dose of whole bacteria contained 5-10-fold more LPS than was presented
previously in purified form (5), the deacylating capacity of AF may
have been exceeded in the current experiments with whole bacteria. In
support of this explanation, increasing the amount of purified LPS
presented to AF decreased the percentage deacylated.2
The addition of PMN during AF-mediated extracellular killing of
bacteria did not enhance deacylation (Fig. 3B), similar to the lack of contribution of added PMN to AF-mediated deacylation of
purified LPS (5). In contrast, MNC caused progressive deacylation of
LPS during AF-mediated extracellular killing of bacteria reaching 40-50% by 20 h (Fig. 3C). Uptake of LPS by the two
cell types during extracellular killing was not assessed,
but because PMN express at least ~5-fold less mCD14 (26) and hence
bind purified LPS less effectively than do MNC (5), PMN are also likely
to internalize less LPS from extracellularly disassembled bacteria than
MNC.
However, when intact bacteria were killed intracellularly after
ingestion, bacterial LPS was deacylated by both MNC and PMN. As shown
by retention of 3-OH-14:0 and loss of NFA in the interface (Fig. 4),
this deacylating activity of the cells was acyloxyacyl hydrolase-like.
Although ingestion of intact bacteria by PMN and MNC was similar, the
rate and extent of LPS deacylation was much greater in MNC (~50% by
4 h versus 20-30% by 20 h in PMN), consistent with a ~5-fold higher acyloxyacyl hydrolase activity in these cells
(5).
The finding that PMN do partially deacylate LPS of ingested bacteria
but do not deacylate purified LPS or LPS shed from bacteria in AF may
be explained by avid phagocytosis of intact bacteria by PMN and limited
ability to internalize cell-free LPS (5). Thus, the physical
presentation of LPS may determine the delivery to different host cells
and, therefore, the extent of their participation in deacylation and
detoxification of LPS.
We have shown before that phagocytosis of E. coli by rabbit
exudate cells is accompanied by extensive degradation of bacterial macromolecules (9, 10). Ingestion of E. coli LCD25 by whole exudate cells (~90% PMN) also triggered extensive degradation of
bacterial RNA (Fig. 5) and hydrolysis of most of the bacterial phospholipids by both phagocyte types (Fig. 4). However, only MNC
deacylated LPS almost as efficiently as phospholipids (Fig. 4B), consistent with a prominent role of MNC in LPS detoxification.
How LPS is detoxified during an infection in vivo is likely
to be influenced prominently by the extent to which the invading bacteria are disassembled by extracellular factors. When invading organisms in the blood stream are relatively fragile and are destroyed extracellularly or when antibiotics are present, neutralization of LPS
by plasma (lipo)proteins may be most important in detoxification. Such
complexing is viewed as an important step in the delivery of LPS to the
liver and its subsequent disposal (24, 27-29). At extravascular sites
where lipoprotein-mediated clearance is less prominent, delivery of
shed LPS to MNC and subsequent deacylation may be more important. In
addition, at localized inflammatory sites (e.g. the
peritoneal cavity), cationic proteins such as the
bactericidal/permeability-increasing protein (BPI) (13) may both
promote ingestion of bacteria (30) leading to deacylation and
detoxification by phagocytes and prevent protracted LPS signaling by
complexing shed LPS (31). The mobilization of extracellular acyloxyacyl
hydrolase at inflammatory sites may contribute further to LPS
detoxification, especially when LPS loads are lower than those
presented in this study. Moreover, because plasma acyloxyacyl hydrolase
levels in the rabbit are markedly elevated in some inflammatory settings (32), extracellular LPS deacylation in vivo may be more prominent than shown here. Inactivation of LPS either by complexing with lipoproteins or by deacylation is not as fast (Refs. 5
and 31 and this study) as the very rapid lipopolysaccharide-binding protein and CD14-mediated LPS signaling and therefore should not interfere with initial activation of host defenses but prevent continued and excessive host responses.
Our findings suggest that the fate of the LPS in Gram-negative bacteria
that are resistant to extracellular disassembly (25, 33) will depend on
intracellular events after ingestion and particularly upon the
participation of MNC. Because PMN greatly outnumber mononuclear
phagocytes, the majority of bacterial invaders are initially ingested
by PMN. However, after intravenous administration of smooth bacteria to
rats, immunologically detectable LPS is found primarily in mononuclear
phagocytes (8). The present study shows that mononuclear phagocytes are
more effective at LPS deacylation than PMN, suggesting eventual
translocation of LPS from PMN to MNC.
Two mechanisms may be proposed for such a transfer. 1) LPS and
partially deacylated species may be released from PMN (34) and
subsequently taken up by mononuclear phagocytes. 2) Bacteria-laden PMN
may become apoptotic (35) and, consequently, subject to ingestion by
macrophages (36, 37) in a process leading to dampening of inflammatory
responses (38).
In summary, deacylation of bacteria-associated or shed LPS over the
course of several hours allows initial signaling of host cells and
up-regulation of antibacterial defenses but may curb prolonged and
potentially self-destructive signaling as LPS remains present at many
sites in the host after infection. This study provides the first
evidence that mammalian phagocytes in a local inflammatory setting can
degrade the endotoxic moiety of LPS when it is presented as a
constituent of whole bacteria.
We thank Dr. Richard Darveau, Dr. Jan Vilcek,
John Gerecitano, and the members of our group for advice and assistance.
*
This work was supported in part by United States Public
Health Service Grants R37 DK 05472 and AI 18188 and by the Xoma Corp. (Berkeley, CA).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
§
Supported by National Institutes of Health Training Grant
5T32GM07308 (NIGMS).
**
To whom correspondence should be addressed: New York University
School of Medicine, Dept. of Microbiology, 550 1st Ave., New York, NY
10016. Tel.: 212-263-5633; Fax: 212-263-8276; E-mail: Elsbap01@mcrcr.med.nyu.edu.
2
Y. Weinrauch, unpublished observations.
The abbreviations used are:
LPS, lipopolysaccharide(s);
12:0, lauric acid;
14:0, myristic acid;
16:0, palmitic acid;
3-OH-14:0, 3-hydroxymyristic acid;
C6d UBAF, unbound
fraction of CM-Sephadex-treated ascitic fluid from C6-deficient
rabbits;
CL, cardiolipin;
HBSS+, Hanks' balanced salts
solution without phenol red;
HBSS
Deacylation of Lipopolysaccharide in Whole Escherichia
coli during Destruction by Cellular and Extracellular Components
of a Rabbit Peritoneal Inflammatory Exudate*
§,,
**, and
Medicine and
Microbiology, New York University School of Medicine, New
York, New York 10016, ¶ Departments of Internal Medicine and
Microbiology, University of Texas Southwestern Medical Center, Dallas,
Texas 75235, and Department of Medicine and
Microbiology, Inflammation Program, University of Iowa School of
Medicine, Iowa City, Iowa 52242
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
) divalent cations,
were from Whittaker Bioproducts, Walkersville, MD). HEMA-3 stain was
from Biochemical Sciences, INC (Swedesboro, NJ). Cycloheximide was from
Sigma. Trichloroacetic acid was from Fisher. CM-Sephadex was from
Amersham Pharmacia Biotech. Glass microfibre (GF/C) filters were from
Whatman (Clifton, NJ). Aquasol-2 scintillation fluid was from Packard
Instrument Co. (Meriden, CT). Silica gel G plates were from Analtech
(Newark, DE); reverse-phase KC18 plates were from Whatman. Radiolabeled
thin-layer chromatography (TLC) standards, [1-14C]12:0
(58 mCi/mmol), n-[9,10-3H]14:0 (53 Ci/mmol),
n-[9,10-3H]16:0, and [1-14C]18:0
(57 mCi/mmol) were all from Amersham Pharmacia Biotech. Unlabeled 18:1,
PE, PG, PC, LPE, and LPG standards were from Sigma. [14C]LPS was purified from Escherichia coli
LCD25 as described previously (11).
containing
0.1% HSA). Resuspended cells were then recombined with the supernatant
fluid, and samples were precipitated for >20 min at 4 °C with 500 µl of ice-cold 20% trichloroacetic acid. Precipitates were poured
over 24-mm circular glass microfibre (GF/C) filters, and tubes and
filtered sediment were washed 3 times with 1.5 ml of 10% ice-cold
trichloroacetic acid. Filters were washed once with ice-cold 95%
ethanol and air-dried. Precipitates were solubilized with 100 µl of
2% SDS and counted in a Beckman LS6000IC liquid scintillation counter
using Aquasol-2 scintillant.
98% of the radiolabeled
lipids in the chloroform phase remained at the origin of
TLC1; 91.1 ± 2.4% (mean ± S.E.) of these
counts were in intact PL. The % of bacterial phospholipids that
remained intact during subsequent incubation was calculated by the
following formula: (% of total cpm in CHCl3 phase) × (% of CHCl3 phase cpm at the origin in
TLC1) × (% of this material comprising PE + PG + CL,
as determined by TLC2)/(% of total cpm in intact bacterial
PL at time = 0).
Partitioning and relative amounts of 3-OH-14:0 in the fatty acids of E. coli LCD25 LPS
Silica gel G plates were developed
for 45-50 min with petroleum ether/diethyl ether/acetic acid (70:30:1;
v:v:v) to separate PL/lyso-PL (at origin), 3-OH 14:0, NFA, and
triglycerides (at solvent front). Samples were run in parallel with
standards: 3-OH-[14C]14:0 (purified by TLC from
hydrolyzed 14C acetate-labeled E. coli or
purified LPS) and [1-14C]12:0.
Silica gel G plates were developed
for 100-110 min with chloroform/methanol/H2O/acetic acid
(65:25:4:1; v:v:v:v) to separate various PL and lyso-PL species.
Migration of unlabeled 18:1, PE, PG, PC LPE, and LPG standards was
determined after detection by iodine vapor staining.
Reverse-phase KC18 plates were
developed twice for 45-50 min each with acetic acid/acetonitrile (1:1;
v:v) to separate individual fatty acids in samples or radiolabeled
standards (n-[9,10-3H]12:0,
(n-[9,10(n)-3H]14:0;
[1-14C]16:0, [1-14C]18:0) loaded in
parallel lanes. Labeled lipid standards were detected by AMBIS and/or
by en3Hance (NEN Life Science Products) autoradiography.
containing 0.1% HSA to remove adherent but
nonphagocytosed bacteria. Radioactivity in the pellet, supernatant, and
washes was measured by scintillation counting. In addition, cell smears
were prepared after 15- and 60-min by cytospin and were stained with
HEMA-3.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
95% PL, FFAs, and other degradation products of PL were
recovered in the chloroform phase (18). Although the IF of extracted
bacteria contained fatty-acylated compounds other than LPS, acyl groups
from these contaminants co-migrated chiefly with 16:0 on TLC (compare
lanes 3 and 6 of Fig. 2B). Evidence that these contaminants contained little or no 12:0 and 14:0 is provided by the closely similar relative amounts of 3-OH-14:0, 12:0,
and 14:0 in the interface of extracted bacteria and purified LPS (Table
IB). Thus, measurement of the loss of 3-OH-14:0, 12:0, and 14:0 from
the IF allowed clear assessment of LPS deacylation.
View larger version (30K):
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Fig. 1.
Schematic representation of procedures used
for lipid analyses.
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[in a new window]
Fig. 2.
TLC of fractions derived from purified LPS
and E. coli labeled with [14C]-acetate.
E. coli LCD25 were labeled with [14C]acetate
during growth as described under "Experimental Procedures." LPS was
prepared from radiolabeled E. coli as described previously
(11). See Fig. 1 for details of preparation of samples for TLC
analysis. A, TLC1. Lane 1, purified
[14C]12:0 and 3-OH-[14C]14:0 standards;
lane 2, hydrolyzed purified LPS; lane 3,
hydrolyzed whole bacteria; lane 4, hydrolyzed interface
material from extracted bacteria; lane 5, hydrolyzed
CHCl3 phase material from extracted bacteria; lane
6, CHCl3 phase from extracted bacteria.
CHCl3 phase of extracted cell suspensions after a 20-h
incubation of bacteria with AF (lane 7) or with C6-deficient
UBAF + PMN (lane 8) or plus MNC (lane 9) is
shown. B, TLC3. Lane 1, purified
[14C]12:0 and [3H]14:0 standards;
lane 2, [14C]12:0, -16:0, and -18:0 standards.
Total NFA recovered from hydrolyzed purified LPS (lane 3),
hydrolyzed bacteria (lane 4), hydrolyzed CHCl3
phase material from extracted bacteria (lane 5), and
hydrolyzed interface material from extracted bacteria (lane
6) is shown. Radiolabeled species in lane 1 of
panel B were detected by fluorography; all other samples
were detected by proportional argon ionization.
97% by 1 h, data not shown). This killing was
accompanied by hydrolysis of >95% bacterial PL by 1 h (Fig.
3A). In contrast, no loss of
3-OH-14:0, 12:0, or 14:0 from bacterial LPS was observed even after
incubation for 20 h (Fig. 3A).
View larger version (15K):
[in a new window]
Fig. 3.
Deacylation of bacterial LPS and PL during
incubation of [14C]acetate-labeled E. coli
LCD25 with 90% AF alone (A) or in the presence
of PMN (B) or MNC (C).
Incubations and assays of the time-dependent loss of 12:0
( 
), 14:0 (
), and 3-OH-14:0 (
) from LPS and loss of intact
bacterial PL (
) were carried out as described under "Experimental
Procedures." The data shown represent the mean (± S.E.) of at least
four separate determinations from at least two separate experiments.
Where error bars are not visible, they are smaller than the plot
symbols.
View larger version (15K):
[in a new window]
Fig. 4.
Deacylation of bacterial LPS and PL during
incubation of [14C]acetate-labeled E. coli
LCD25 with PMN (A) or MNC (B)
in C6-deficient UBAF. See the legend to Fig. 3. Incubations and
assays of the time-dependent loss of 12:0 ( ), 14:0
(), and 3-OH-14:0 () from LPS and loss of intact bacterial PL
() were carried out as described under "Experimental
Procedures."
85%
PMN). In AF alone, degradation of RNA was maximal at 2 h, reaching
70%, whereas protein degradation reached a plateau of 10% by 1 h
(Fig. 5A). Ingestion by
exudate leukocytes was accompanied by rapid degradation of RNA that
reached its maximum of 65-75% by 2 h, whereas protein
degradation reached a plateau of ~30% by 1 h (Fig.
5B). Thus, like PL, degradation of RNA and proteins in
E. coli LCD25 is rapid, reaching an early plateau.
Degradation of RNA by both cellular and extracellular components
of the rabbit exudate was particularly extensive, as has been shown
previously in different bacterial species (9, 10, 19). Therefore,
deacylation of LPS by AF and PMN is slow and limited relative
to degradation of other bacterial components.
View larger version (12K):
[in a new window]
Fig. 5.
Time-dependent degradation of
biosynthetically labeled E. coli LCD25 RNA and
proteins during killing by rabbit AF and inflammatory peritoneal
exudate cells. Incubations and measurements of protein ( 
) and
RNA (
) degradation were carried out as described under
"Experimental Procedures." A, rabbit AF; B,
peritoneal exudate leukocytes in 50% C6-deficient UBAF. These data
represent one (RNA) or the mean ± range of two (protein)
determinations. TCA, trichloroacetic acid.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
ACKNOWLEDGEMENTS
FOOTNOTES
ABBREVIATIONS
, Hanks' balanced salts
solution without phenol red, Ca2+, and
Mg2+;
HSA, human serum albumin;
IF, interface
in Bligh-Dyer extraction between CHCl3 and
CH3OH/H2O phases;
LPE, lysophosphatidylethanolamine;
LPG, lysophosphatidylglycerol;
MNC, mononuclear cells;
FA, fatty acids;
NFA, nonhydroxylated FA;
FFA, free
FA;
PE, phosphatidylethanolamine;
PG, phosphatidylglycerol;
PMN, polymorphonuclear leukocytes;
PL, phospholipids;
AF, ascitic
fluid.
REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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