Neutrophil elastase and syndecan shedding contribute to ...

R E S E A R C H A R T I C L E

Neutrophil elastaseand syndecan shedding contribute toantithrombindepletion inmurineanthraxMyung-Chul Chung, Shelley C. Jorgensen, Taissia G. Popova, Charles L. Bailey & Serguei G. Popov

National Center for Biodefense and Infectious Diseases, George Mason University, Manassas, VA, USA

Correspondence: Serguei G. Popov,

National Center for Biodefense and Infectious

Diseases, George Mason University, 10900

University Blvd., Manassas, VA 20110, USA.

Tel.: 11 703 993 4713; fax: 11 703 993

4288; e-mail: [email protected]

Received 29 May 2008; revised 25 August

2008; accepted 26 August 2008.

First published online 8 October 2008.

DOI:10.1111/j.1574-695X.2008.00480.x

Editor: Nicholas Carbonetti

Keywords

anthrax; disseminated intravascular

coagulation; sepsis; antithrombin; syndecan.

Abstract

Bacillus anthracis infection is associated with severe hemostatic disturbances but

their roles and contribution to fatality remain incompletely characterized. We

undertook analyses of circulating antithrombin levels during the course of infection

using a comparison of lethal and nonlethal murine anthrax models. Plasma samples

were obtained from DBA/2 mice challenged intraperitoneally with the spores of

either toxigenic B. anthracis Sterne strain or nontoxigenic, avirulent delta Sterne

strain. We found that plasma antithrombin levels were rapidly depleted in Sterne

spore-challenged mice, concomitant with elevation of neutrophil elastase (NE) and

massive syndecan shedding from the liver into circulation. The shed syndecan

bound with antithrombin accelerated NE-mediated antithrombin proteolysis. The

liver response to infection demonstrated strain-specific compensatory increases of

antithrombin and syndecan gene transcription. Both bacterial strains induced

changes in blood coagulation parameters consistent with the onset of disseminated

intravascular coagulation. We propose that antithrombin depletion proceeding

through activation of neutrophils and massive shedding of heparin-like syndecan

from the liver into circulation contribute to anthrax coagulopathy.

Introduction

Bacillus anthracis is a gram-positive, spore-forming bacter-

ium. It is the causative agent of anthrax – a highly dangerous

disease of humans and several animal species. The disease

can be manifested in one of the three distinct forms, namely

cutaneous, gastrointestinal, and inhalational, depending on

the route of the host exposure to the infectious bacterial

spores. If left untreated, anthrax can quickly progress

systemically, which is associated with high mortality. Several

pathogenic factors of B. anthracis have been identified and

characterized. It is commonly accepted that lethal and

edema toxins (LT and ET, respectively), along with the

antiphagocytic capsule are the major virulence factors of

the bacterium (Prince, 2003; Moayeri & Leppla, 2004;

Heninger et al., 2006). Recently, other secreted bacterial

proteins, including broad-spectrum proteases and hemo-

lysins, have been reported as substantial contributors to

anthrax pathology and mortality (Popov et al., 2005; Mosser

& Rest, 2006; Heffernan et al., 2007). Although studies on

the effects of B. anthracis pathogenic factors in vivo and in

vitro have yielded important insights into their function, the

pathological mechanisms underlying high mortality of

systemic anthrax, as well as the immediate cause of death in

this disease remains incompletely understood.

Hemostatic and hemodynamic abnormalities during an-

thrax infection are well known (Smith & Keppie, 1954;

Abramova et al., 1993; Guarner et al., 2003; Stearns-Kurosawa

et al., 2006). Dilated capillaries are such a common morpho-

logic feature of anthrax infection that for many years before

the discovery of anthrax toxins, the mechanical obstruction

of capillaries by the bacilli was believed to be the cause of

death in anthrax septicemia (Albrink, 1961). This theory

was dismissed based on the fact that anthrax-infected

animals at times die in the absence of severe bacteremia

(Eckert & Bonventre, 1963). However, later experiments

revealed that widespread pulmonary capillary and kidney

glomeruli thrombosis in infected rats, rabbits, and guinea-

pigs was not caused solely by the mechanical obstruction of

capillaries by a large number of bacilli (Dalldorf & Beal,

1967). The authors suggested thrombosis as the major cause

of death and concluded that anthrax pathogenic factors

acted directly upon the membranes of capillary endothelium

to cause increased permeability and capillary thrombosis.

FEMS Immunol Med Microbiol 54 (2008) 309–318 c� 2008 George Mason UniversityNational Center for Biodefense

Journal compilation c� 2008 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd.

Dow

nloaded from https://academ

ic.oup.com/fem

spd/article/54/3/309/512730 by guest on 04 July 2022

Further studies (Grinberg et al., 2001; Culley et al., 2005;

Stearns-Kurosawa et al., 2006) strongly supported these

observations, and it has been recognized recently (Berry

et al., 2003) that vascular pathology in anthrax patients is

indicative of the onset of condition known as disseminated

intravascular coagulation (DIC) or consumptive coagulo-

pathy (Stearns-Kurosawa et al., 2006).

DIC is often associated with microbial sepsis and typically

sets off intravascular thrombosis, tissue hypoxia, activation

of the fibrinolytic pathways, depletion of coagulation fac-

tors, inability to maintain hemostasis, and overt bleeding

(Opal & Esmon, 2003). All these features are present in the

course of anthrax infection, and some of them, such as

thrombocytopenia and vascular leakage, have also been

reported in the cell and animal models of anthrax toxemia

(Gozes et al., 2006; Stearns-Kurosawa et al., 2006). Clinical

manifestations of DIC may be underlined by different

disorders and range from the absence of overt symptoms to

shock and death. The pathogenic mechanisms that trigger

DIC in anthrax are currently unknown, and the contribu-

tion of DIC to anthrax mortality remains to be elucidated.

Available evidence indicates that the coagulopathic effect of

the anthrax infectious process is complex, and likely repre-

sents a concerted action of toxins (Ascenzi et al., 2002;

Moayeri & Leppla, 2004; Popova et al., 2006; Turk, 2007;

Watson et al., 2007) and other pathogenic factors such as

broad-spectrum proteases and hemolytic proteins (Bonventre

& Eckert, 1963; Popov et al., 2005; Chung et al., 2006;

Heffernan et al., 2007). We recently reported that

B. anthracis-challenged mice demonstrate increases in the

amount of circulating soluble proteoglycans, namely synde-

can (SDC)-1, -4, as well as plasma depletion of the func-

tional von Willebrand factor and its natural regulator

ADAMTS13, implicating these pathogenic processes in the

development of hemorrhage (Popova et al., 2006; Chung

et al., 2008). Bacillus anthracis proteases have been shown to

be hemorrhagic in the mouse skin test (Popov et al., 2005),

and able to degrade a range of proteins critically involved in

the regulation of hemostasis in vitro, such as a2-macroglo-

bulin, a2-antiplasmin, and a1-protease inhibitor, which are

the most important serum regulators of plasmin and blood

elastase (Chung et al., 2006). With regard to blood fibrino-

lysis, B. anthracis proteases can also cleave fibrinogen and

therefore may impair the clotting process during anthrax

infection (Chung et al., 2006, 2008).

In sepsis, antithrombin is quickly depleted by virtue of its

consumption during complex formation with activated

clotting factors and after proteolytic degradation by elastase

secreted from polymorphonuclear leukocytes (Jordan et al.,

1987). Antithrombin is a plasma-derived serine protease

inhibitor from a single chain glycoprotein, which controls

the activity of thrombin and the inhibition of several other

proteases of the coagulation cascade. Antithrombin has

potent anticoagulant activity and is significantly enhanced

by heparin and vessel wall-associated glycosaminoglycans

(GAGs). Despite acting as an acceleratory cofactor for

antithrombin, heparin was also found to considerably accel-

erate the in vitro inactivation of antithrombin by neutrophil

elastase (NE) (Jordan et al., 1987). Sharp decreases of

antithrombin in sepsis has been shown previously to cause

a dramatic imbalance in hemostasis and homeostasis result-

ing in an excess of activated factors. In fact, when anti-

thrombin levels decrease to o 30% of normal, DIC patients

show almost absolute lethality with sepsis (Roemisch et al.,

2002). Although B. anthracis infection induces DIC with

severe sepsis, the underlying mechanisms responsible

remain poorly understood.

In the present study, we undertook analyses of circulating

antithrombin levels and several other hemostatic parameters

in the murine model of anthrax. Plasma samples were

obtained from mice challenged with the spores of either

toxigenic B. anthracis Sterne strain (pXO11, pXO2�), which

is virulent in DBA/2 mice, or nontoxigenic, avirulent delta

Sterne strain (pXO1�, pXO2�). Strong antithrombin deple-

tion from circulation at the late stages of toxigenic infection

was observed compared with nontoxigenic one. By bio-

chemical analyses, we demonstrate that activation of neu-

trophils and massive shedding of heparin-like syndecan

from the liver into circulation is one of the pathological

mechanisms of antithrombin depletion.

Materials and methods

Animals

Female, 9-week-old DBA/2 mice were purchased from

Jackson Laboratory (Bar Harbor, ME) and maintained at

Biocon Inc. (Rockville, MD). Food and water were supplied

ad libitum. Animals were allowed to acclimate to their

surroundings for 1 week before experiments. The George

Mason University Institutional Animal Care and Use Com-

mittee and Biocon Animal Care and Use Committee/Insti-

tutional Review Board have approved all protocols before

animal experiments.

Preparation of mouse plasma

Mice (n = 5–10 per treatment group) were challenged in-

traperitoneally (i.p.) with 5� 106 spores of B. anthracis

nonencapsulated Sterne strain 34F2 (pXO11, pXO2�) or

nontoxigenic delta Sterne strain (pXO1�, pXO2�) obtained

from the Colorado Serum Company (Boulder, CO) and

National Center for Biodefense and Infectious Diseases

(Manassas, VA), respectively. Approximately 200–500 mL of

blood were drawn from the survived animals (n = 3–5 per

treatment group at every time point) via the retro-orbital

FEMS Immunol Med Microbiol 54 (2008) 309–318c� 2008 George Mason UniversityNational Center for BiodefenseJournal compilation c� 2008 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd.

310 M.-C. Chung et al.

Dow


ic.oup.com/fem


sinus into 50 mL of 0.5 M EDTA and centrifuged at 900 g for

10 min to obtain platelet poor plasma (PPP).

Bacterial load and course of infection

Mice were euthanized at 24, 48, and 72 h post-spore

challenge and the livers were collected to measure bacterial

dissemination. Liver tissues were homogenized in

phosphate-buffered saline (PBS), serially diluted, and plated

on Luria–Bertani agar to determine numbers of CFU. The

highest bacterial burden was 2.0� 105 CFUg�1 (48 h) in liver

of Sterne-challenged mice and 5.2� 104 CFUg�1 (72 h) in

delta Sterne-challenged mice. The 50% mortality took place

at 72 h post-Sterne challenge; there was 100% survival in all

delta Sterne-spore challenged mice (Supporting Informa-

tion, Fig. S1).

Heparin cofactor antithrombin assay

For the quantitative determination of antithrombin in

mouse plasma, a heparin cofactor assay kit (Chromogenix,

Coamatic Antithrombin Kit) was used according to the

manufacturer’s recommendation. Briefly, 50mL of 1 : 20

diluted plasma were mixed with 50 mL of human factor Xa

in the presence of heparin and incubated at 37 1C for 90 s in

a 96-well microtiter plate. To measure the residual factor Xa

activity, 50 mL of substrate (N-a-Cbo-Arg-Gly-Arg-p-

nitroanilide) were added to the reaction for 90 s. Then 50mL

of 20% acetic acid were added to each microplate well and

the absorbance was read at 405 nm. Relative antithrombin

activity (%) was evaluated based on a standard curve using

pooled control plasma.

Western blot

Plasma samples (corresponding to 20 mg of total protein)

from normal control and spore-challenged mice were boiled

in sodium dodecyl sulfate (SDS) sample buffer containing

30 mM dithiothreitol. The samples were loaded onto a

NuPAGE 4–12% Bis Tris Gel (Invitrogen) and transferred

onto a polyvinylidene difluoride membrane using the iBlot

Gel Transfer System (Invitrogen). The membrane was

blocked with a solution of 5% milk powder in PBS,

incubated with primary sheep a-mouse antithrombin anti-

body (Abcam) or mouse a-human NE antibody (Santa Cruz

Biotechnology), and finally incubated with an horse radish

peroxidase (HRP)-conjugated secondary antibody (rabbit

a-sheep IgG from Abcam, or sheep a-mouse IgG from GE

Healthcare, correspondingly). The membranes were devel-

oped using an ECL Western blotting detection kit (Amer-

sham Biosciences) and Kodak BioMax light film or a

Molecular Imager ChemiDoc XRS System (Bio-Rad).

Ligand blot

Two micrograms of human antithrombin (Sigma) were

loaded onto a NuPAGE 4–12% Bis-Tris Gel (Invitrogen)

and transferred onto a nitrocellulose membrane using the

iBlot Gel Transfer System (Invitrogen). The membrane was

incubated overnight at 4 1C in PBS containing 0.05%

Tween-20 (PBST), washed three times with PBST, incubated

with the protein ligand solution in PBST for 1.5 h at room

temperature and then incubated with a corresponding

primary antibody (1 : 3000 dilution) for 2 h. Primary anti-

bodies were sheep anti-mouse antithrombin (Abcam),

mouse anti-mouse NE (Santa Cruz Biotechnology), mouse

anti-mouse SDC-1 (BD Pharmingen), goat anti-mouse

SDC-4 (Santa Cruz Biotechnology) and chicken anti-

human a1-protease inhibitor (Abcam). Culture supernatants

of anthrolysin O (ALO)-treated normal murine mammary

gland cells (NMuMGCs) (150mL) were prepared by treating

the cultured cells with 1 mg mL�1 of ALO for 24 h to induce

syndecan shedding as described previously (Popova et al.,

2006). Finally, the membranes were washed and detected

with corresponding secondary antibodies described above.

Immunoprecipitation

Pooled plasma (30mL) from 48-h Sterne spore-challenged

mice was diluted in PBS (1 : 10) and precleared by adding

20 mL of suspended (25% v/v) protein A/G-agarose beads

(Santa Cruz Biotechnology). Primary antibodies (2mg)

against antithrombin, NE, or SDC-1 were added after

incubation for 1 h at 4 1C. Then, 20 mL of protein A/G-

agarose beads were added followed by incubation at 4 1C on

a platform rocker overnight. The immunoprecipitates were

collected by centrifugation at 900 g for 30 s at 4 1C and

washed four times with 1.0 mL of PBS. After the final wash,

the proteins were eluted by directly adding 40mL of 2�electrophoresis sample buffer. After boiling for 3 min, the

samples were subjected to electrophoresis and Western blot

analysis with antithrombin antibody as described above.

Enzyme-linked immunosorbent assays (ELISAs)

Wells of a 96-mictotiter (NUNC MaxiSorp) plate were

coated with 50 mL of plasma samples from five mice

(0.01–20mg of total protein) in 50 mM carbonate buffer

(pH 9.0) and incubated overnight at 4 1C. Coated wells were

blocked with 1% bovine serum albumin in PBS, and

primary antibodies were added in a 1 : 3000 dilution. The

antibodies were against the following antigens: mouse

a-human D-dimer antibody (Abcam, ab-10050); goat

a-mouse PAI-1 (Santa Cruz Biotechnology, sc-6644);

goat a-mouse PAI-2 (Santa Cruz Biotechnology, sc-6649);

rabbit a-mouse a2-antiplasmin (Abcam, ab-28326); rabbit

a-mouse plasminogen (Abcam, ab-28319); mouse a-human



311Antithrombin depletion in a murine anthrax model

Dow


ic.oup.com/fem


thrombin (Abcam, ab-17199); sheep a-human antithrom-

bin (Abcam, ab-8776); and chicken a-human a1-antitrypsin

(Abcam, ab-14226). All antibodies against human proteins

cross-reacted with corresponding mouse proteins. The

pooled plasma samples from five mice were incubated in

triplicates in the coated wells at room temperature for 1 h,

and washed three times with PBST. Fifty microliters of

corresponding HRP-conjugated secondary antitotal IgG

antibodies were added in a 1 : 10 000 dilution followed by

1-h incubation at room temperature. After washing five

times with PBST, the tetramethylbenzidine (TMB) substrate

(100mL) was added. The reaction was stopped with 1 M

H2SO4 (50 mL) and the absorbance was read at 450 nm.

Standard curves were made from a dilution of normal

mouse plasma and the results were expressed relative to

controls. For TAT analysis, wells of a 96-well plate (NUNC

MaxiSorp) were coated with 100 mL of anti-antithrombin

antibody (1 mg mL�1 in coating buffer). Plates were incu-

bated overnight at 4 1C. The plates were washed three times

with PBST, and 200mL of PBS/10% fetal calf serum (FCS)

was added to each well as a blocking step for 1 h followed by

addition of 50 mL of standard or fourfold-diluted plasma

samples in PBST/1% FCS for 1 h on a shaker. The plate was

washed three times with PBST and then incubated with

100mL of antithrombin antibody (0.5mg mL�1) in PBST/1%

FCS for 1 h. The wells were washed three times with PBST

and incubated with 100 mL of HRP-antibody in PBST

followed by washes and TMB solution as described above.

Reverse-transcriptase (RT)-PCR

For determination of SDC-1, -4, and antithrombin mRNA

expression, total RNA was prepared from frozen liver

sections stored at � 80 1C. Random-primed cDNA was

prepared from 5mg of total RNA using the Platinum PCR

Supermix (Invitrogen). Specific primers for antithrombin,

50-ATG GTA CAT TCC TCA CCC TGC G (sense) and 50-

AGC ACC ACT GCT CCC ACA ATG A (antisense); SDC-1,

50-ATG AGA CGC GCG GCG CTC TGG CTT (sense) and

50-GGCGTA GAA CTC CTC CTG CTT GGT (antisense);

SDC-4, 50-AAC CAC ATC CCT GAG AAT GC (sense) and

50-AGG AAA ACG GCA AAG AGG AT (antisense) were

used during RT-PCR. After amplification, PCR products

were separated on a 1.7% agarose gel and stained with

ethidium bromide. The gel photographs were analyzed by

densitometry and the expression levels were normalized by

the amount of b-actin mRNA.

Statistical analyses

A paired two-tailed Student’s t-test was used for all statistical

analyses. The results are presented as means� SD. Values of

Po 0.05 were considered statistically significant.

Results

Depletion of antithrombin in plasma isassociated with its proteolytic degradation

Reduced antithrombin activity in the blood is often con-

sidered one of the most important diagnostic indicators of

nonsurvival in sepsis (Spero et al., 1980; Bick et al., 1996). In

order to examine circulating antithrombin levels, DBA/2

mice were challenged intraperitoneally with 5� 106 spores

of toxigenic Sterne or nontoxigenic delta Sterne strains.

Plasma samples from randomly chosen individual mice were

prepared using EDTA, loaded onto the SDS-polyacrylamide

gel electrophoresis (PAGE) and analyzed using Western blot.

Sterne-challenged mice demonstrated a strong reduction in

the overall intensity of bands recognized by the antithrom-

bin-specific antibody (Fig. 1a and c). The reduction was

most intense between 24 and 48 h postchallenge, while

animals sacrificed for analysis at 72 h postchallenge demon-

strated less pronounced antithrombin depletion. For inter-

pretation of these results, it has to be taken into account that

before 48 h postchallenge the population of animals is

characteristic of the disease onset and is not yet skewed by

high mortality, which takes place around 72 h (Popov et al.,

2005). In contrast, at 72 h postchallenge the live animals

represent mainly resistant survivors. In comparison, plas-

mas from delta Sterne-challenged mice showed only subtle

changes in the intensities of the bands immunoreacting with

the anti-antithrombin antibody (Fig. 1b and c). At 72 h

post-delta Sterne-challenge all mice survived, and the

amount of antithrombin in plasma returned to its level in

healthy mice (Fig. 1c) indicating convalescence after disease.

Next, we used the heparin cofactor assay to test the

amount of functional antithrombin in the plasma of

Sterne-challenged mice. This assay is based on the ability of

antithrombin to inhibit the activity of factor Xa in the

presence of heparin. The assay is able to discriminate

between antithrombin-deficient and antithrombin nondefi-

cient individuals better than thrombin-based assays

(Demers et al., 1993) and gives accurate results in patients

receiving heparin therapy (Bohner et al., 1994). We found

that the heparin cofactor activity gradually decreased after

Sterne spore challenge (Fig. 1d). At 72 h postinfection, the

heparin cofactor activity remained low, indicating a func-

tional deficiency of circulating antithrombin in spite of the

fact that the animals survived up to this time showed total

antithrombin protein levels close to normal (Fig. 1a and c).

In order to determine whether the depletion of antith-

rombin from circulation could be explained by decreased

expression of the antithrombin gene, total RNA was pre-

pared from the liver of B. anthracis spore-challenged mice

was analyzed using RT-PCR. The results showed that during

the course of toxigenic infection, the levels of antithrombin



Dow


ic.oup.com/fem


gene expression steadily increased, while the nontoxigenic

infection did not induce a statistically significant response

(Fig. 2). This suggested that depletion of antithrombin in

plasma was associated with its proteolytic degradation, but

not suppression of the antithrombin gene expression.

Shed SDC-1 and -4 interact with antithrombinand accelerate neutrophil elastase-mediatedantithrombin proteolysis in vitro

NE expression significantly correlates with DIC severity and

has a good predictive value for mortality in patients

suspected of having DIC (Song et al., 2007). During DIC,

NE is able to inactivate antithrombin by a specific and

limited proteolytic cleavage (Jordan et al., 1987). We, there-

fore, suggested that antithrombin depletion found in toxi-

genic B. anthracis-challenged mice could be relevant to the

induction of NE during infection and tested with NE

antibodies the plasma samples previously used for the

analysis of antithrombin. NE levels were significantly in-

creased at 24 and 48 h post-Sterne spore challenge (Fig. 3).

Western blot analysis of antithrombin and NE from plasma

of each mouse demonstrated that high levels of NE corre-

lated with the reduced levels of antithrombin (Figs 1 and 3).

To prove a possible causal connection between enzymatic

activity of NE and antithrombin levels, we carried out

experiments with purified NE. Inactivation of antithrombin

by NE is known to be considerably accelerated by heparan

sulfate proteoglycans derived from endothelial cells (Jordan

et al., 1987). On the surface of endothelium, antithrombin

associates with proteoglycans such as SDC-1 and -4 posses-

sing sulfated carbohydrate chains structurally similar to

heparin, and the anticoagulant heparin subfraction also

demonstrates high-affinity binding for antithrombin

(Jordan et al., 1987; San Antonio et al., 1994; Woods et al.,

1998, 2000; Ueno et al., 2001). Bacillus anthracis accelerates

SDC-1 shedding due to the activity of proteolytic and

hemolytic proteins (Popova et al., 2006). Therefore, we

tested whether shed syndecans accelerate NE-dependent

antithrombin proteolysis. First, we investigated the interac-

tion of isolated antithrombin and NE proteins with shed

syndecans using the ligand blot assay. As a source of

syndecans, we used culture supernatant of NMuMGCs

treated with B. anthracis hemolysin ALO (Popova et al.,

Fig. 1. Antithrombin protein level and functional activity are reduced in plasma of spore-challenged mice. Western blots of antithrombin in plasma from

Sterne (a) and delta Sterne (b) spore-challenged mice. Numbers indicate individual mice for comparison with NE analysis in Fig. 3. Arrows indicate the positions

on gels corresponding to molecular mass of antithrombin. (c) Densitometric analysis of antithrombin bands in (a) and (b). Open and gray bars indicate relative

antithrombin levels in plasmas from delta Sterne- and Sterne-challenged mice, correspondingly. �P = 0.001 and ��P = 0.025 relative to control mice. (d)

Heparin cofactor activity of antithrombin in plasma of Sterne spore-challenged mice relative to unchallenged control. �P = 0.02 and ��Po 0.01 compared

with control unchallenged mice.

Fig. 2. Gene expression for antithrombin measured using RT-PCR rela-

tive to actin in spore-challenged mouse plasma. Three mice from each

group were subjected to RT-PCR. Expression was calculated as a fold

change relative to control (mean� SD). P-values were evaluated by

paired Student’s t-test.




Dow


ic.oup.com/fem


2006). Antithrombin blotted onto a nitrocellulose mem-

brane was incubated with NE, culture supernatant, or

a1-protease inhibitor as a negative control. After intensive

washing, interaction of antithrombin with the overlaid

proteins was detected using specific antibodies. The results

demonstrated that antithrombin interacted with NE, shed

SDC-1 and -4 from culture supernatant, but not with

a1-protease inhibitor (Fig. 4a). The interaction between

antithrombin and NE or syndecan was further confirmed

using immunoprecipitation analysis (Fig. 4b).

Bacillus anthracis -challenged mice showsyndecan shedding in liver blood vessels

Damage to the liver plays a central role in abnormal

hemostasis and the development of DIC (Mammen, 1992).

Syndecan shedding can be stimulated by different cell stress

factors including B. anthracis-secreted proteins, and the liver

has been suggested as one of the primary targets (Watchorn

et al., 2002; Kovalszky et al., 2004; Popova et al., 2006). Using

immunofluorescence microscopy with a SDC-1-specific

primary antibody and a fluorescein-labeled secondary anti-

body, we evaluated the amount of syndecan in the liver of

spore-challenged mice. Control samples demonstrated that

SDC-1 was readily detectable as green fluorescence in the

lining of normal liver sinusoids and the lumen of liver blood

vessel was stained negative (Fig. 5a and c). Conversely,

Sterne spore-challenged mice demonstrated a considerable

increase in the amount of expressed SDC-1 and shortening

of hepatocyte cords indicating a physiological response of

the liver to infection (Fig. 5b). Concomitantly, a massive

amount of SDC-1-positive aggregates was detected in the

lumen of blood vessels (Fig. 5d) in agreement with our

previous ELISA data demonstrating the appearance of shed

SDC-1 in the blood of infected mice (Popova et al., 2006).

SDC-1 gene expression was increased in both Sterne and

delta Sterne-challenged mice (Fig. 6a), which has been

shown previously to correlate with changes in cytoskeleton

organization (Lebakken & Rapraeger, 1996). SDC-4-

deficient mice have impaired repair to tissue injury and are

more susceptible to thrombus formation and endotoxin

shock (Ishiguro et al., 2001). In our study, SDC-4 gene

expression was upregulated only by the toxigenic strain

(Fig. 6b).

Markers for coagulopathy in plasma ofB. anthracis spore-challenged mice

In order to further characterize hemostatic abnormalities

during anthrax infection we analyzed plasmas of challenged

mice using a conventional ELISA with specific antibodies

against several key proteins associated with DIC (Table 1).

According to the current understanding of DIC, the deter-

mination of fibrin degradation products in plasma is critical

for diagnosis (Levi & ten Cate, 1999; Schrecengost et al.,

2003). Analysis of D-dimers, the breakdown products of the

fibrin mesh, helps to differentiate degradation of cross-

linked fibrin formed in the process of coagulation from

fibrinogen degradation products. Bacteremia typically re-

sults in a rapidly occurring increase in fibrinolytic activity,

which becomes impaired due to a sustained level of

Fig. 4. Antithrombin interacts with human NE, shed SDC-1 and SDC-4

in vitro and in vivo. (a) Ligand blot analysis. Antithrombin was immobi-

lized on the nitrocellulose membrane, incubated with indicated analytes,

and immunodetected using specific antibodies. a1-PI is shown as a

negative control. Culture supernatant of ALO-treated NMuMGCs served

as a source of SDC-1. (b) Immunoprecipitation (IP) of proteins from

plasma of Sterne spore-challenged mice at 24 and 48 h postinfection

with antithrombin-, NE-, and SDC-1-specific antibodies demonstrate the

presence of bound antithrombin detected using immunoblotting (IB)

with antithrombin-specific antibody.

Fig. 3. NE level is increased in plasma of Sterne spore-challenged mice.

(a) Western blot of NE in plasma of Sterne spore-challenged mice. The

major band with a molecular mass around 60 kDa in unchallenged

plasma corresponds to NE. Open triangles indicate complexes of NE with

unidentified proteins detected with anti-NE antibody during infection.

Numbers indicate individual mice for comparison with antithrombin

analysis in Fig. 1a. (b) Densitometric analysis of NE bands in (a).



Dow


ic.oup.com/fem


plasminogen activator inhibitor-1 (PAI-1), the main regu-

lator of endogenous fibrinolysis (Paul et al., 2005). In

agreement with this, Table 1 shows that progression of the

infectious process in the case of both B. anthracis strains

resulted in a sharp increase in D-dimer activity as early as

24 h postinfection followed by a slow decline at 72 h. The D-

dimer formation was accompanied by the generation of PAI-

1, and to the lesser extent, of PAI-2. This process took place

faster in the case of the virulent infection (Po 0.05 at 24 h).

In agreement with moderate elevation of plasmin–antiplasmin

complex (PAP) in DIC patients (Mavrommatis et al., 2001),

the levels of PAP in our experiments were slightly increased

above normal indicating that the generation of plasmin and

the fibrinolytic process were impaired.

The dynamics of DIC can be judged by measuring plasma

concentration of thrombin–antithrombin (TAT) complexes.

This marker is highly sensitive to low-grade activation of

coagulation but may have a limited specificity in complex

pathological conditions because it reflects the net effect of

generation and consumption of both thrombin and antith-

rombin. Our results demonstrate that TAT levels gradually

increased and became significantly elevated (Po 0.05) at

72 h postinfection in both Sterne and delta Sterne spore-

challenged mice. Taken together with the depletion of the

levels of circulating antithrombin (Fig. 1), these data in-

dicate a quick onset of a prothrombotic condition during

anthrax infection.

Discussion

The host septic response plays a key role in anthrax

pathogenesis (Smith & Keppie, 1954; Stearns-Kurosawa

et al., 2006). Bacterial infections, in particular septicemia,

are the most common causes of DIC (Kreger et al., 1980;

Mavrommatis et al., 2001). However, in anthrax, DIC as an

underlying pathophysiological condition attracted little

Fig. 5. Immunofluorescent staining of liver

using SDC-1-specific primary antibody and

fluorescein-labeled secondary antibody. SDC-1

(green) in the lining of sinusoids (a, b) and in the

lumen of liver blood vessel of control (a, c) and

Sterne spore-challenged mice at 72 h postinfection

(p.i.) (b, d). Brightness of images in (c) and (d) has

been increased to demonstrate accumulation

of SDC-1 aggregates in vessel lumen (d).

Fig. 6. Gene expression for SDC-1 (a), and SDC-4 (b) measured using

RT-PCR relative to actin in spore-challenged mouse plasma. Three mice

from each group were subjected to RT-PCR. Expression was calculated as

a fold change relative to control. P-values were calculated using paired

Student’s t-test.




Dow


ic.oup.com/fem


attention, although it has been identified in victims of the

1979 Sverdlovsk, Russia accident (Abramova et al., 1993;

Meselson et al., 1994) and the 2001 bioterror attacks in the

United States (Berry et al., 2003; Guarner et al., 2003). DIC is

a disorder with highly variable clinical presentations, making

straightforward clinical and laboratory diagnosis compli-

cated. No single laboratory test is sensitive or specific enough

to allow a definite diagnosis. Intrinsic variability between

patients and the rarity of human inhalational anthrax cases

present additional complications and emphasize the value of

inbred animal models. Although previous experimental

animal studies present features of B. anthracis infection

consistent with DIC (Stearns-Kurosawa et al., 2006), only a

limited number of blood coagulation parameters have been

tested. In this study, we expanded characterization of DIC in

the murine anthrax model and directly compared plasma

from animals challenged with lethal and nonlethal anthrax

strains. We found that infection with both B. anthracis

strains elicits similar changes of several coagulation markers

indicative of the septic response to systemic infection, but

lethally challenged animals demonstrate an aggravated re-

sponse incompatible with survival. Our findings support the

opinion that severe infection or sepsis is almost invariably

associated with activation of coagulation without major

differences in causative microorganisms (Levi & van der

Poll, 2004). It is the intensity of coagulation and inflamma-

tion responses that spans from a subtle prohemostatic

response in delta Sterne-challenged mice to fully activated

DIC and subsequent death in Sterne-challenged ones.

Our data are consistent with the current view on the

pathogenesis of DIC, in which impaired fibrinolysis is

mediated by the release of plasminogen activators from

endothelial cells immediately followed by an increase in the

plasma levels of PAI-1. On the other hand, fibrin formation

caused by tissue factor-mediated generation of thrombin is

accompanied by dysfunction of inhibitory mechanisms,

such as the antithrombin system (Franchini et al., 2006;

Levi, 2007). Typically, depletion of antithrombin is a result

of increased consumption, enzymatic degradation, impaired

liver synthesis and vascular leakage (Franchini et al., 2006).

The broad spectrum of antithrombin’s capacity to inhibit

almost all coagulation factors renders it the most important

serpin in hemostasis. A decrease in antithrombin below 70%

is considered to be a highly significant imbalance in hemo-

stasis and homeostasis resulting in an excess of activated

factors and the proinflammatory condition often resulting

in death (Roemisch et al., 2002). Complete deficiency of

antithrombin induces embryonic lethality in antithrombin-

deficient mice with extensive subcutaneous hemorrhage

(Ishiguro et al., 2000). We demonstrate that both lethal and

nonlethal B. anthracis challenges induced increased levels of

circulating TAT consistent with consumption of antithrom-

bin in inactive complex forms. Our data also demonstrate

that the toxigenic spore challenge was associated with a

strong decrease in the levels of antithrombin (down to about

30% of its normal level according to Western blot assay) and

correspondingly elevated levels of NE, similar to plasma of

patients with DIC or organ failure (Wada, 2004). Invasion of

neutrophils also correlates with poor outcome in anthrax

patients (Abramova et al., 1993). Although our animal

model was matched by inbred genetic background, gender

and age, we found a remarkable variation in the magnitude

of individual animal responses suggesting that differences in

the immune status of individual mice might be relevant to

the intrinsic variability of DIC symptoms.

In the case of both B. anthracis strains the depletion of

antithrombin in our animal model was not caused by

suppression of antithrombin gene expression in the liver.

Table 1. Markers for coagulopathy in plasma from Sterne (S) and delta Sterne (dS) spore-challenged mice

Marker Challenge strain

Time postchallenge (h)

0 24 48 72

D-dimer S 100� 35 1260�397�� 1410� 470�� 830� 614�

dS 1002�432�� 1187� 471�� 986� 179��

PAI-1 S 100� 39 299�55� 329� 213� 300� 93�

dS 151�25�� 350� 142� 180� 35�

PAI-2 S 100� 36 125�18 157� 26 181� 37

dS 112�55 115� 15 151� 50

PAP S 100� 15 112�39 101� 20 139� 70

dS 83�29 126� 55 168� 41�

TAT S 100� 75 66�46 121� 50 232� 109

dS 116�105 71� 24 203� 58�

a1-PI S 100� 31 143�17 109� 55 71� 15

dS 151�27 67� 33 112� 45

ELISA values are calculated as a percent to control (mean� SD) for plasma samples from five different mice.�Po 0.05, ��Po 0.01, ��Po 0.001 compared with the PBS-treated control mice (paired Student’s t-test, n = 5).



Dow


ic.oup.com/fem


Antithrombin is known to be susceptible to NE-mediated

cleavage in the presence of blood proteoglycans, such as

SDC-1 (Jordan et al., 1987). We reported previously in-

creased shedding of syndecans into the blood of B. anthracis

spore-challenged mice (Popova et al., 2006), and our present

results implicate syndecan shedding as the cause of acceler-

ated antithrombin degradation by NE. However, activation

of NE may not be the only mechanism of antithrombin

depletion during infection. Relevant to this, vascular leakage

and hemorrhage have been demonstrated previously in mice

administered LT and other secreted proteases (Cui et al.,

2005; Culley et al., 2005).

In summary, our findings reveal early aggressive hemo-

static changes in the murine anthrax model consistent with

DIC, acceleration of the coagulation cascade, and subse-

quent host septic response, which need to be taken into

account in future therapeutic interventions.

Acknowledgement

This work was supported by the United States Department

of Defense grant DAMD 17-03-C-01220.

Authors’contribution

M.-C.C. and S.C.J. contributed equally to this study.

References

Abramova F, Grinberg L, Yampolskaya O & Walker D (1993)

Pathology of inhalational anthrax in 42 cases from the

Sverdlovsk outbreak of 1979. Proc Natl Acad Sci USA 90:

2291–2294.

Albrink W (1961) Pathogenesis of inhalation anthrax. Bacteriol

Rev 25: 268–273.

Ascenzi P, Visca P, Ippolito G, Spallarossa A, Bolognesi M &

Montecucco C (2002) Anthrax toxin: a tripartite lethal

combination. FEBS Lett 531: 384–388.

Berry K, Colvin S, Blythe D et al. (2003) Follow-up of deaths

among U.S. postal service workers potentially exposed to

Bacillus anthracis – district of Columbia, 2001–2002. JAMA

290: 2119–2120.

Bick R, Strauss J & Frenkel E (1996) Thrombosis and hemorrhage

in oncology patients. Hematol Oncol Clin North Am 10:

875–907.

Bohner J, von Pape K & Blaurock M (1994) Thrombin-based

antithrombin assays show overestimation of antithrombin III

activity in patients on heparin therapy due to heparin cofactor

II influence. Thromb Haemost 71: 280–283.

Bonventre P & Eckert N (1963) The biologic activities of Bacillus

anthracis and Bacillus cereus culture filtrates. Am J Pathol 43:

201–211.

Chung MC, Popova TG, Millis BA, Mukherjee DV, Zhou W,

Liotta LA, Petricoin EF, Chandhoke V, Bailey C & Popov SG

(2006) Secreted neutral metalloproteases of Bacillus anthracis

as candidate pathogenic factors. J Biol Chem 281:

31408–31418.

Chung MC, Popova TG, Jorgensen SC, Dong L, Chandhoke V,

Bailey CL & Popov SG (2008) Degradation of circulating von

Willebrand factor and its regulator ADAMTS13 implicates

secreted Bacillus anthracis metalloproteases in anthrax

consumptive coagulopathy. J Biol Chem 283: 9531–9542.

Cui X, Li Y, Moayeri M et al. (2005) Late treatment with a

protective antigen-directed monoclonal antibody improves

hemodynamic function and survival in a lethal toxin-infused

rat model of anthrax sepsis. J Infect Dis 191: 422–434.

Culley N, Pinson D, Chakrabarty A, Mayo M & Levine S (2005)

Pathophysiological manifestations in mice exposed to anthrax

lethal toxin. Infect Immun 73: 7006–7010.

Dalldorf F & Beal F (1967) Capillary thrombosis as a cause of

death in experimental anthrax. Arch Path 83: 154–161.

Demers C, Henderson P, Blajchman M et al. (1993) An

antithrombin III assay based on factor Xa inhibition provides a

more reliable test to identify congenital antithrombin III

deficiency than an assay based on thrombin inhibition.

Thromb Haemost 69: 231–235.

Eckert N & Bonventre P (1963) In vivo effects of Bacillus anthracis

culture filtrates. J Infect Dis 112: 226–232.

Franchini M, Lippi G & Manzato F (2006) Recent acquisitions in

the pathophysiology, diagnosis and treatment of disseminated

intravascular coagulation. Thrombosis J 4: 4.

Gozes Y, Moayeri M, Wiggins J & Leppla S (2006) Anthrax lethal

toxin induces ketotifen-sensitive intradermal vascular leakage

in certain inbred mice. Infect Immun 74: 1266–1272.

Grinberg R, Abramova F, Yampolskaya O, Walker D & Smith J

(2001) Quantitative pathology of inhalational anthrax I:

quantitative microscopic findings. Mod Pathol 5: 482–495.

Guarner J, Jernigan J, Shieh W, Tatti K, Flannagan L & Stephens D

(2003) Pathology and pathogenesis of bioterrorism-related

inhalational anthrax. Am J Path 163: 701–709.

Heffernan B, Thomason B, Herring-Palmer A & Hanna P (2007)

Bacillus anthracis anthrolysin O and three phospholipases C

are functionally redundant in a murine model of inhalation

anthrax. FEMS Microbiol Lett 271: 98–105.

Heninger S, Drysdale M, Lovchik J, Hutt J, Lipscomb M &

Koehler T (2006) Toxin-deficient mutants of B. anthracis are

lethal in a murine model for pulmonary anthrax. Infect Immun

74: 6067–6074.

Ishiguro K, Kojima T, Kadomatsu K et al. (2000) Complete

antithrombin deficiency in mice results in embryonic lethality.

J Clin Invest 106: 873–878.

Ishiguro K, Kadomatsu K, Kojima T et al. (2001) Syndecan-4

deficiency leads to high mortality of lipopolysaccharide-

injected mice. J Biol Chem 276: 47483–47488.

Jordan R, Kilpatrick J & Nelson R (1987) Heparin promotes the

inactivation of AT by neutrophil elastase. Science 14: 777–779.

Kovalszky I, Dudas J, Gallai M, Hollosi P, Tatrai P, Tatrai E &

Schaff Z (2004) Proteoglycans in the liver. Magy Onkol 48:

207–213.




Dow


ic.oup.com/fem


Kreger B, Craven D & McCabe W (1980) Gram-negative

bacteremia. IV. Re-evaluation of clinical features and

treatment in 612 patients. Am J Med 68: 344–355.

Lebakken C & Rapraeger A (1996) Syndecan-1 mediates cell

spreading in transfected human lymphoblastoid (Raji) cells.

J Cell Biol 132: 1209–1221.

Levi M (2007) Disseminated intravascular coagulation. Crit Care

Med 35: 2191–2195.

Levi M & ten Cate H (1999) Disseminated intravascular

coagulation. N Engl J Med 341: 586–592.

Levi M & van der Poll T (2004) Coagulation in sepsis: all bugs bite

equally. Crit Care Clin 8: 99–100.

Mammen E (1992) Coagulation abnormalities in liver disease.

Hematol Oncol Clin North Am 6: 1247–1257.

Mavrommatis AC, Theodoridis T, Economou M, Kotanidou A, El

Ali M, Christopoulou-Kokkinou V & Zakynthinos SG (2001)

Activation of the fibrinolytic system and utilization of the

coagulation inhibitors in sepsis: comparison with severe sepsis

and septic shock. Intensive Care Med 27: 1853–1859.

Meselson M, Guillemin J, Hugh-Jones M, Langmuir A, Popova I,

Shelokov A & Yampolskaya O (1994) The Sverdlovsk anthrax

outbreak of 1979. Science 266: 1202–1208.

Moayeri M & Leppla S (2004) The roles of anthrax toxin in

pathogenesis. Curr Opin Microbiol 7: 19–24.

Mosser E & Rest R (2006) The Bacillus anthracis cholesterol-

dependent cytolysin, Anthrolysin O, kills human neutrophils,

monocytes and macrophages. BMC Microbiol 6: 56.

Opal S & Esmon C (2003) Bench-to-bedside review: functional

relationships between coagulation and the innate immune

response and their respective roles in the pathogenesis of

sepsis. Crit Care 7: 23–38.

Paul R, Winkler F, Bayerlein I, Popp B, Pfister H & Koedel U

(2005) Urokinase-type plasminogen activator receptor

regulates leukocyte recruitment during experimental

pneumococcal meningitis. J Infect Dis 191: 776–782.

Popov SG, Popova TG, Hopkins S, Weinstein RS, MacAfee R,

Fryxell KJ, Chandhoke V, Bailey C & Alibek K (2005) Effective

antiprotease-antibiotic treatment of experimental anthrax.

BMC Infect Dis 5: 25–39.

Popova TG, Millis B, Bradburne C, Nazarenko S, Bailey C,

Chandhoke V & Popov SG (2006) Acceleration of epithelial

cell syndecan-1 shedding by anthrax hemolytic virulence

factors. BMC Microbiol 6: 8.

Prince A (2003) The host response to anthrax lethal toxin:

unexpected observations. J Clin Invest 112: 656–658.

Roemisch J, Gray E, Hoffmann JN & Wiedermann CJ (2002)

Antithrombin: a new look at the actions of a serine protease

inhibitor. Blood Coagul Fibrinolysis 13: 657–670.

San Antonio JD, Karnovsky MJ, Gay S, Sanderson RD & Lander

AD (1994) Interactions of syndecan-1 and heparin with

human collagens. Glycobiology 4: 327–332.

Schrecengost JE, LeGallo RD, Boyd JC, Moons KG, Gonias SL,

Rose CE Jr & Bruns DE (2003) Comparison of diagnostic

accuracies in outpatients and hospitalized patients of D-dimer

testing for the evaluation of suspected pulmonary embolism.

Clin Chem 49: 1483–1490.

Smith H & Keppie J (1954) Observations on experimental

anthrax: demonstration of a specific lethal factor produced in

vivo by B. anthracis. Nature 173: 869–870.

Song S, Kim H, Park M & Cho H (2007) Neutrophil CD64

expression is associated with severity and prognosis of

disseminated intravascular coagulation. Thromb Res 121:

499–507.

Spero J, Lewis J & Hasiba U (1980) Disseminated intravascular

coagulation. Findings in 346 patients. Thromb Haemost 43:

28–33.

Stearns-Kurosawa D, Lupu F, Taylor F, Kinasewitz G & Kurosawa S

(2006) Sepsis and pathophysiology of anthrax in a nonhuman

primate model. Am J Pathol 169: 433–444.

Turk B (2007) Manipulation of host signalling pathways by

anthrax toxins. Biochem J 402: 405–417.

Ueno M, Yamada S, Zako M, Bernfield M & Sugahara K (2001)

Structural characterization of heparin sulfate and chondroitin

sulfate of syndecan-1 purified from normal murine mammary

gland epithelial cells. J Biol Chem 276: 29134–29140.

Wada H (2004) Disseminated intravascular coagulation. Clin

Chim Acta 344: 13–21.

Watchorn TM, Waddell I & Ross JA (2002) Proteolysis-inducing

factor differentially influences transcriptional regulation in

endothelial subtypes. Am J Physiol Endocrinol Metab 282:

E763–E769.

Watson L, Mock J, Lal H et al. (2007) Lethal and edema toxins of

anthrax induce distinct hemodynamic dysfunction. Front

Biosci 1: 4670–4675.

Woods A & Couchman J (1998) Syndecans: synergistic activators

of cell adhesion. Trends Cell Biol 8: 189–192.

Woods A, Longely R, Tumova S & Couchman J (2000) Syndecan-

4 binding to the high affinity heparin-binding domain of

fibronectin drives focal adhesion formation in fibroblasts. Arch

Biochem Biophys 374: 66–72.

Supporting Information

Additional Supporting Information may be found in the

online version of this article:

Fig. S1. Survival curve of DBA/2 mice (n = 10) challenged

intraperitoneally (i.p.) with 5� 106 spores of Bacillus an-

thracis Sterne strain 34F2 (open circles) or nontoxigenic

delta Sterne strain (closed circles).

Please note: Wiley-Blackwell is not responsible for the

content or functionality of any supporting materials sup-

plied by the authors. Any queries (other than missing

material) should be directed to the corresponding author

for the article.



Dow


ic.oup.com/fem


Neutrophil elastase and syndecan shedding contribute to ...

Documents

Transcript of Neutrophil elastase and syndecan shedding contribute to ...