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Cell Motility and the Cytoskeleton 40:71–86 (1998)
FtsZ From Escherichia coli, Azotobacter
vinelandii, and Thermotoga
GTP Hydrolysis, and Assembly
Chunlin Lu, Jesse Stricker, and Harold P. Erickson*
Department of Cell Biology, Duke University Medical Center,
Durham, North Carolina
We have cloned the ftsZ genes from Thermotoga maritima and Azotobacter
vinelandii and expressed the proteins (TmFtsZ and AzFtsZ) in Escherichia coli.
We compared these proteins to E. coli FtsZ (EcFtsZ), and found that several
remarkable features of their GTPase activities were similar for all three species,
implying that these characteristics may be universal among FtsZs. Using a
calibrated protein assay, we found that all three FtsZs bound 1 mole guanine
nucleotide per mole FtsZ and hydrolyzed GTP at high rates (.2 GTP per FtsZ per
min). All three required magnesium and a monovalent cation for GTP hydrolysis.
Previous reports showed that EcFtsZ (and some other species) required potassium.
We confirmed this specificity for EcFtsZ but found that potassium and sodium both
worked for Az- and TmFtsZ. Specific GTPase activity had a striking dependence
on FtsZ concentration: activity (per FtsZ molecule) was absent or low below 50
µg/ml, rose steeply from 50 to 300 µg/ml and plateaued at a constant high value
above 300 µg/ml. This finding suggests that the active state requires a polymer that
is assembled cooperatively at 50–300 µg/ml. A good candidate for the active
polymer was visualized by negative stain electron microscopy—straight protofilaments and protofilament pairs were seen under all conditions with active GTPase.
We suggest that the GTP hydrolysis of FtsZ may be coupled to assembly, as it is for
tubulin, with hydrolysis occurring shortly after an FtsZ monomer associates onto a
protofilament end. As a part of this study, we determined the concentration of
EcFtsZ and TmFtsZ by quantitative amino acid analysis and used this to
standardize the bicinchonic acid colorimetric assay. This is the first accurate
determination of FtsZ concentration. Using this standard and quantitative Western
blotting, we determined that the average E. coli cell has 15,000 molecules of FtsZ,
at a concentration of 400 µg/ml. This is just above the plateau for full GTPase
activity in vitro. Cell Motil. Cytoskeleton 40:71-86, 1998. r 1998 Wiley-Liss, Inc.
Key words: tubulin; dynamic instability; nucleation; cell division; bicinchonic acid; quantitative Western
The bacterial cell division protein FtsZ has been
found in more than three dozen prokaryotic species,
including archaebacteria, and in chloroplasts [Erickson,
1997]. It is a major cytoskeletal protein of bacteria,
forming a ring that constricts at the site of septation
[Addinall et al., 1996; Ma et al., 1996]. FtsZ is a
homologue of tubulin, as demonstrated first by sequence
r 1998 Wiley-Liss, Inc.
Contract grant sponsor: National Institutes of Health; Contract grant
number: GM28553
*Correspondence to: Harold P. Erickson, Department of Cell Biology,
Duke University Medical Center, Durham, NC 27710;
Received 26 November 1997; accepted 21 January 1998
Lu et al.
alignment [Mukherjee and Lutkenhaus, 1994], and recently confirmed by the close similarity of the atomic
structures [Löwe and Amos, 1998; Nogales et al., 1998].
The homology is functional as well, as Escherichia coli
FtsZ (EcFtsZ) can assemble in vitro into protofilament
sheets with a lattice similar to that of the microtubule
wall [Erickson et al., 1996]. FtsZ protofilaments can also
adopt a sharply curved conformation, forming minirings
very similar to tubulin rings [Erickson et al., 1996;
Erickson and Stoffler, 1996]. The structural homology of
FtsZ and tubulin polymers suggests that the cytoskeletal
function of these proteins has been conserved from
bacterial FtsZ to eukaryotic tubulins [Erickson et al.,
1996; Erickson, 1997].
Both tubulin and FtsZ bind and hydrolyze GTP. The
GTPase activity of tubulin is tightly coupled to assembly.
In the absence of assembly the rate of hydrolysis is very
low, but after a subunit enters the microtubule lattice its
bound GTP is rapidly hydrolyzed [O’Brien et al., 1987].
EcFtsZ purified from overexpression in bacteria has
demonstrated a very high level of GTPase [RayChaudhuri and Park, 1992; de Boer et al., 1992; Mukherjee et
al., 1993].
In the present study we have reinvestigated and
extended the characterization of the GTPase of EcFtsZ.
As a part of this study, we have calibrated the assay for
FtsZ protein concentration and determined the concentration in E. coli. In addition, we have cloned the ftsZ genes
from Azotobacter vinelandii and Thermotoga maritima,
expressed these proteins in E. coli (AzFtsZ and TmFtsZ),
and compared the GTPase and polymer state of the FtsZ
from the three species.
Cloning AzFtsZ and TmFtsZ
Two degenerate oligonucleotide primers were designed based on the most highly conserved regions,
NTDNQA and GGGTGTG of the N-terminal portion of
FtsZ (forward primer, 58-AA(A/T)AC(G/A/T/C)GA(C/T)GC(G/A/T/C)CA(A/G)GC-38 reverse primer, 58-GT(G/A/
C)CC-38). Genomic DNA from Thermotoga maritima was
generously provided by Dr. Michael Adams (University
of Georgia, Athens, GA). Polymerase chain reaction
(PCR) products were separated on an agarose gel, and
fragments in the range of 200–500 base pairs (bp) were
isolated and cloned. A 350-bp fragment was identified
that showed significant homology to ftsZ of other species.
Thermotoga genomic DNA was cut with EcoRI and
circularized, and inverse PCR was performed as described by Ochman and Hartl [1988] with erratic but
eventually successful results. Each attempt at inverse
PCR gave many clones with unrelated sequence, but
some showed homology to ftsZ of other species, extending the sequence in the 58 and 38 directions. New primers
were designed near the limits of these extended sequences, and the inverse PCR was repeated. Sequences
were eventually found that extended past the initiation
and stop codons, as judged by homology with ftsZ
sequences of other species. The final sequence was
obtained from the pET-FtsZ plasmid (described below),
which was produced with a single PCR reaction from
genomic DNA using the error-correcting Pfu polymerase.
The same PCR primers were used to amplify
genomic DNA of Azotobacter vinelandii (strain CA,
provided by Dr. Paul E. Bishop, North Carolina State
University, Raleigh). A 350-bp PCR product was then
used to screen a cosmid library, provided by Dr. Limin
Zhang (Virginia Polytechnic Institute, Blacksburg, VA).
A positive cosmid was identified and digested by EcoRI.
A ,7-kbp fragment that hybridized with the PCR probe
was gel-purified and subcloned into the SuperCos 1
cosmid vector (Stratagene, LaJolla, CA). The ftsZ gene was
sequenced in both directions starting from the region identified by PCR, and the full-length ftsZ was obtained as judged
by comparison to the ftsZ sequences of other species.
Construction of Recombinant Expression Vector
for TmFtsZ and AzFtsZ
Once we had identified the initiation and stop
sequences of Thermotoga ftsZ, we designed primers to
amplify the full-length coding sequence, adding an NdeI
site at the 58 end and a BamHI sequence at the 38 end.
PCR amplification used 2.5 units of Pfu DNA polymerase
(Stratagene, LaJolla, CA) in a 50-µl reaction mixture
containing 0.2 mM dNTP, 1 µM primers, 0.5 µg Thermotoga genomic DNA and the buffer recommended by
Stratagene. The predicted 1,070-bp fragment was isolated
from an agarose gel, digested with NdeI and BamHI at
37°C for 24 h, gel purified again, and then ligated into
NdeI and BamHI sites of expression vector pET-11b
(Novagen, Madison, WI). Both strands of the resulting
recombinant vector pET-TmFtsZ were sequenced from
the synthetic NdeI site through the BamHI site to determine
the final version of the Thermotoga ftsZ sequence.
The same procedure was used to produce a pET-11
expression vector for AzFtsZ. The pET expression plasmid for EcFtsZ was provided by Dr. David Bramhill,
Merck [Bramhill and Thompson, 1994] and was also
duplicated in our laboratory.
Expression and Purification of Ec-, Az-,
and TmFtsZ
EcFtsZ, AzFtsZ, and TmFtsZ were successfully
overproduced in E. coli strain BL21(DE3) transformed
with pET-11 expression vectors containing the coding
region of the three genes. Cultures for all three were
GTPase and Assembly of FtsZ
grown in LB medium at 37°C to A600 ,1.2 and induced
by adding 0.25–0.5 mM IPTG. We also induced some
cultures at A600 ,0.7 and found a slightly lower yield of
FtsZ. The protein appeared identical at the two levels of
expression; in particular the ratio of protein precipitated
at 20% and 25% ammonium sulfate (see below). After a
3-h induction, bacteria from a 1-L culture were centrifuged and suspended in 20 ml resuspension buffer (0.05
M Tris, 0.1 M NaCl, 1 mM EDTA, 1 mM PMSF, pH 8.0).
Lysozyme was added to 0.4 mg/ml, and the mix was
incubated on ice for 2 h; MgCl2 was added to 5 mM and
the sample was frozen overnight at 220°C. The bacteria
were thawed and sonicated twice for 30 s on ice. DNase I
was added to 10 µg/ml and incubated 60 min at 4°C. Cell
walls and insoluble debris was removed by centrifugation. All three FtsZ proteins were soluble in E. coli
cytoplasm, and represented more than one-half of the
total soluble protein.
In earlier preparations of FtsZ, we added Triton
X-100 to 1% after 30 min with lysozyme, continued
agitation for 5 min and froze the mixture. However, the
triton caused the ammonium sulfate precipitate to partially float when centrifuged, instead of forming a compact pellet. This was a problem especially for TmFtsZ
whose precipitation required higher ammonium sulfate.
We have subsequently eliminated Triton from the lysis
buffer and have found that freezing and sonication
efficiently lyses the bacteria.
After centrifuging the lysed bacteria, EcFtsZ was
precipitated from the bacterial supernatant with ammonium sulfate. Initially we used 20% saturated ammonium
sulfate, which precipitated virtually all the FtsZ and
almost no bacterial proteins in the presence of 1% Triton.
After eliminating Triton we found that less than one-half
of the EcFtsZ was precipitated at 20% saturated ammonium sulfate, and it had a substantially lower GTPase
activity than protein precipitated at 25%. We then modified our purification protocol. We first did a cut at 20%
saturated ammonium sulfate, discarding the pellet (about
30–40% of the FtsZ). We then precipitated the remaining
EcFtsZ by raising the ammonium sulfate to 25% saturation. The pellet was resuspended in one-half the original
volume of resuspension buffer supplemented with 10 mM
Mg and re-precipitated with 25% saturated ammonium
sulfate. At this stage, EcFtsZ appeared pure on sodium
dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE), and we found that ion-exchange chromatography
on DEAE Sephacel or mono-Q did not result in any
apparent increase in purity. For the experiments reported
in this paper, EcFtsZ was prepared by the 20–25%
ammonium sulfate precipitation, with no additional steps.
TmFtsZ required 38% saturated ammonium sulfate
for precipitation, and many host proteins were pelleted
with it. To separate contaminating proteins from TmFtsZ,
a heating step was introduced to denature and precipitate
the E. coli proteins. We found that TmFtsZ was substantially stabilized when 1 mM GTP and 5 mM Mg were
added before heating. With GTP and Mg, TmFtsZ
remained soluble after 20 min at 70°C or 10 min at 80°C,
while most contaminating proteins precipitated. For purification, a 2- to 3-ml sample of TmFtsZ in a centrifuge
tube was heated to 80°C for 2 min, cooled on ice for 20
min, and then centrifuged. As a final purification step, the
TmFtsZ was chromatographed on a mono-Q column,
where it eluted at 0.37 M NaCl.
AzFtsZ was purified by two cycles of precipitation
at 20% saturated ammonium sulfate. Eventually, all FtsZ
proteins were resuspended in the resuspension buffer with
10 mM Mg, aliquoted, and stored at 280°C.
Calibration of Protein Concentration
Quantitative amino acid analysis of bovine serum
albumin (BSA), EcFtsZ, and TmFtsZ was performed by
Dr. Ida Thorgersen (Macromolecular Structure Facility,
Duke University Medical Center). BSA concentration
was also checked by ultraviolet (UV) spectroscopy. These
calibrated standards were used to calibrate the bicinchonic acid (BCA) colorimetric assay (Pierce Biochemicals, Rockford, IL) [Smith et al., 1986].
Antibody Production
Rabbits were inoculated with 200 µg of GST-FtsZ
fusion protein in 2 ml saline mixed with 2 ml complete
Freund’s adjuvant. One rabbit (FtsZd-5434) was immunized with a protein band that was cut out from an SDS
gel (,2 ml gel was equilibrated in saline and emulsified
with Freund’s adjuvant). Another rabbit (FtsZn-5435)
was injected with soluble, native protein that had been
purified by ammonium sulfate precipitation and affinity
chromatography on a glutathione column. Four boosts of
50 µg protein in 2 ml saline plus 2 ml incomplete
Freund’s adjuvant were given over 10 weeks. All bleeds
for both antisera stained a single band, of varying
intensities, at ,40 kDa, in Western blots of whole E. coli.
Bleed 4 of the FtsZn-5435 antiserum was further purified
by affinity binding to EcFtsZ coupled to CNBr-Sepharose. A total of 1.44 mg of antibody was purified from
5 ml antiserum.
Quantitative Western Blotting
Escherichia coli BL21 cells were used for determination of normal FtsZ levels. Cells were grown in LB at
37°C, shaken at 250 rpm, to an OD600 of 0.5–0.7. Cells
were then killed with 0.02% sodium azide and put on ice
for 30 min. They were then counted with a hemocytometer under 4003 magnification using phase contrast
optics. Several cultures were also diluted and plated on
LB-agar plates before fixing so as to carry out viable
Lu et al.
counts. In each case, viable counts agreed with hemocytometer counts within 10%.
Samples containing a known number of cells were
lysed in SDS buffer and separated on 12% SDSpolyacrylamide reducing gels, transferred to Immobilon
membrane (Millipore, Bedford, MA), and blocked with
5% skim milk/TTBS (100 mM Tris pH 7.4, 150 mM
NaCl, 0.1% Tween) overnight. Primary incubation was
with 0.23 µg/ml FtsZn-5435 affinity purified polyclonal
antibody in 0.25% gelatin/TTBS for 120min. Secondary
incubation was with 2 µCi/blot 125I-labeled protein A
(ICN, Costa Mesa, CA) in 0.25% gelatin/TTBS for
45min. Blots were washed and placed on phosphoimager
screens (Hypercassette, Amersham, Arlington Heights,
IL) for periods ranging from 3 h to 16 h, then scanned and
digitized with a phosphoimager (Fujix Bas1000). The
intensity of lanes on these blots was determined with the
software program MacBas 1.0 (Fuji).
For quantitative Western blotting, blots were run in
sets to ensure consistent results. Three identical blots
containing a set of standards and a set of unknowns and a
blot containing two sets of standards were run concurrently. The dual standards blot allowed confirmation of
consistency within blots, showing that all areas on the
blot were equally sensitive. The multiple blots containing
unknowns allowed confirmation of consistency between
blots. Any set of blots where identical lanes on the dual
standards blot had intensities differing by more than 10%
was discarded. Similarly, in the case in which the
unknowns blots gave intensities differing by more than
10%, they would be discarded. If two of the three
unknowns blots agreed with each other, only the third was
discarded. If all three blots disagreed, all three were
GTP Hydrolysis
FtsZ was desalted, and its buffer was exchanged
using prepacked Econo-Pac 10DG columns (Bio-Rad,
Hercules, CA) in 50 mM Tris-HCl, pH 7.5. Three reaction
buffers were used. Most assays were done in TKM7.5 (50
mM Tris-HCl, pH 7.5, 50 mM KCl, 5 mM Mg acetate).
The second buffer was PB7.7 (physiological buffer: 35
mM MOPS, pH 7.7, 80 mM Glutamate, 350 mM
potassium acetate, 2.5 mM Mg acetate, 50 mM trehalose,
2 mM putrescine), which mimics the ionic composition of
the E. coli cytoplasm [Cayley et al., 1991]. We found that
glutamate, trehalose and putrescine had no effect on
GTPase or assembly, so this buffer is very similar to
TKM7.5, except for the higher potassium concentration.
MEMK6.5 (100 mM MES, pH 6.5, adjusted with KOH, 1
mM EGTA, 5 mM Mg acetate) was a lower pH buffer
used for some assembly and GTPase assays. Buffers
normally contained 0.5 mM GTP (except as noted), and
0.5 µCi (a-32P) GTP (3,000 Ci/mmol). GTP from Amer-
sham was uniformly of high quality when analyzed by
thin layer chromatography, and was used for all assays;
several lots obtained from ICN contained only GDP.
Additives or alterations in the buffer are indicated in the
text. Reaction buffer for TmFtsZ was prewarmed to the
reaction temperature and the reaction was initiated by
adding prewarmed FtsZ to give the indicated concentration. At indicated intervals, 1 µl was withdrawn and
applied to polyethyleneimine cellulose thin layer chromatography plates (10 3 10 cm, Polygramt CEL 300
PEI/UV254, Macherey-Nagel, Germany). The spots dried
within 1–2 min, which we assumed arrested the GTPase.
Plates were developed in 0.75 M KH2PO4, pH 3.4, dried,
and exposed to a phosphoimager plate (IP, FUJIX) for
10–30 min. The imager plate was screened and data were
analyzed using MacBAS1000 v 1.01 software (FUJIX,
BAS 1000Mac, Fuji Photo Film). To avoid errors due to
evaporation of sample during the reaction or imprecision
in the amount of sample loaded, the data were always
read as the ratio of GDP/GTP; this ratio, multiplied by the
known amount of GTP in the starting solution, gave the
amount of GDP produced, which was eventually expressed as moles GDP produced per mole FtsZ.
Because we do not understand the mechanism for
the GTPase reaction, and because variable results have
been reported by different labs, we need to specify the
conditions of our assay precisely. The FtsZ was transferred by gel filtration to 50 mM Tris-HCl, pH 7.5
(containing no GTP, Mg, or K), and frozen at a concentration of 1–3 mg/ml. FtsZ was thawed and kept on ice
before the experiment. A reaction mixture was prepared
containing Tris, KCl, Mg, and GTP to give the desired
final concentrations when mixed with FtsZ in a 20-µl
reaction mixture. FtsZ was added to the reaction mixture
on ice, and the Eppendorf tube was transferred immediately to a 37°C water bath (for Ec- and AzFtsZ). For
TmFtsZ, the concentrated FtsZ and the reaction buffer
containing GTP were separately prewarmed to 70°C and
mixed at the initial time point. Attempts to prewarm
EcFtsZ gave erratic results, perhaps because of denaturation of the protein at 37°C in the absence of GTP.
Assembly of FtsZ
Three buffers were used for assembly studies:
TKM7.5, PB7.7, and MEMK6.5 . FtsZ was transferred to
assembly buffer using 10-ml prepacked Econo-Pac 10DG
columns (Bio-Rad) before assembly. Assembly of TmFtsZ
in MEMK6.5 was started by adding GTP to 2 mM and
incubating at 70°C. At 5 min after incubation at 70°C,
glutaraldehyde (ultrapure TEM grade, Tousimis Research
Corporation, Rockville, MD) was added to 1% to fix the
polymers. The reaction was kept at 70°C for another 2 min,
then cooled to room temperature, and negatively stained
specimens were prepared. EcFtsZ (1 mg/ml) was assembled
GTPase and Assembly of FtsZ
in both MEMK6.5 and PB7.7, while AzFtsZ was polymerized in PB7.7. These reactions were started by adding DEAE
dextran (approximate MW 2,000,000, D-5876, Sigma, St.
Louis, MO) to 0.6 mg/ml and GTP to 2 mM, and incubated on
ice for 10 min, followed by incubation at 37°C for 5 min.
EcFtsZ polymers in the absence of DEAE dextran were
visualized by negative staining at 0°C and 37°C.
Electron Microscopy
Before rotary shadowing, FtsZ was sedimented
through a 5-ml linear gradient of 15–40% glycerol in 0.1
mM GTP, 0.2 M ammonium bicarbonate. Peak fractions
identified by SDS-PAGE were diluted in 40% glycerol,
0.2 M ammonium bicarbonate, sprayed onto freshly
cleaved mica, and rotary shadowed with platinum [Fowler
and Erickson, 1979].
Assembly of FtsZ in vitro was analyzed primarily
by negative stain electron microscopy. Grids coated with
a thin carbon film were subjected to glow discharge to
render the surface hydrophilic and wettable. A total of 10
µl of the assembled FtsZ was applied to the grid,
removed, and drained with filter paper, and 3 drops of
uranyl acetate (a 2% aqueous solution, unadjusted pH,
filtered through 0.2 µm) were washed over the grid while
it was held at a 45° angle. The grid was finally drained
with filter paper and stored. Grids were examined with a
Philips 301 electron microscope and photographed at
Sequences of AzFtsZ and TmFtsZ
and Expression in E. coli
The DNA and deduced amino acid sequences of
AzFtsZ and TmFtsZ have been submitted to the GenBank
database, accession numbers U65939 and U65944. The
amino acid sequence of AzFtsZ is 85% and 83% identical
to FtsZ from the closely related Pseudomonas aeruginosa
and Pseudomonas putida, and 57% identical to EcFtsZ.
TmFtsZ is 45% identical to FtsZ from the gram-positive
bacteria Bacillus subtilis, Staphylococcus aureus, and
Anabaena sp. It is about 40–42% identical to FtsZ from
most gram-negative bacteria (41% to EcFtsZ) and 33–
41% identical to FtsZ from archaebacteria. On a phylogenetic tree the TmFtsZ sequence branches close to the
separation of gram-positive, gram-negative, and archaebacteria [Erickson, 1997]. The 351-amino acid sequence
of TmFtsZ is one of the shortest of 35 known complete
sequences; the only shorter are FtsZ from Haloferax
volcanii [Wang and Lutkenhaus, 1996], and the second,
probably nonessential FtsZ of Rhizobium meliloti [Margolin and Long, 1994].
All three FtsZ proteins were abundantly expressed
as soluble proteins in E. coli. Overexpression of TmFtsZ
did not cause filamentation or otherwise alter cell growth
or viability either before or after induction by IPTG,
suggesting that this protein is inactive in E. coli at 37°C.
However, overexpression of EcFtsZ or AzFtsZ caused
filamentation. Even without induction by IPTG, primary
transformants carrying the EcFtsZ or AzFtsZ recombinant expression plasmid frequently produced smooth
filamentous bacteria, apparently due to the small amount
of expression from the pET vector. Cell filamentation
disappeared after several passages for EcFtsZ-transformed cells, consistent with previous observations [Dai
and Lutkenhaus, 1992]. However, the AzFtsZ-transformed cells remained filamentous, suggesting that even
a small amount of AzFtsZ inhibits the E. coli division
system. This toxicity has been observed for expression of
almost all foreign FtsZ in E. coli [Beall et al., 1988;
Margolin et al., 1991].
Two Fractions of EcFtsZ With Different
GTPase Activities
In our earlier experiments, EcFtsZ was purified by
two cycles of precipitation with 20% saturated ammonium sulfate. However, further experiments suggested
that this EcFtsZ was a mixture of inactive protein and a
fully active form. This preparation represented only
30–40% of the total expressed EcFtsZ; it bound only 0.25
moles of guanine nucleotide per mole of EcFtsZ; and it
had a lower GTPase activity than any other FtsZ preparation. By contrast, the fraction of EcFtsZ precipitated by
25% ammonium sulfate, following a 20% ammonium
sulfate cut, bound 1 mole guanine nucleotide per mole
FtsZ and had a much higher rate of GTP hydrolysis. The
20% cut hydrolyzed 0.31 moles of GTP per mole of
protein per min, compared with 2.4 moles/min for the
25% cut under the same conditions (see below). We
assume that the 25% fraction contained fully active
EcFtsZ and used this for all subsequent experiments.
To check for possible covalent differences in the
two protein preparations, EcFtsZ fractions precipitated at
20% and 25% ammonium sulfate were analyzed by
electrospray mass spectrometry. The measured mass was
40,326.9 Da and 40,327.4 Da, for the two preparations,
identical to the mass of 40327.4 calculated from the
amino acid sequence by the DNAstar (Madison, WI)
program. We conclude that neither preparation of EcFtsZ
has any post-translational modification. Both fractions
also gave identical results upon gradient sedimentation
and electron microscopy (see Fig. 5 for micrographs of
the 25% fraction), suggesting no difference in polymerization state.
Before dismissing the 20% cut as completely uninteresting, we should point out that this sample would
assemble into protofilament sheets in MEMK6.5 and
PB7.7 buffer; remarkably, this assembly occurred without
Lu et al.
addition of DEAE dextran. A maximum of 25–30% of the
protein in the 20% ammonium sulfate fraction could be
pelleted after assembly, compared to 95% of the protein
in the 25% ammonium sulfate fraction, but assembly of
this later fraction was completely dependent on added
DEAE dextran (see below). It is possible that something
in the 20% cut is substituting for DEAE dextran.
Concentration of FtsZ In Vitro and In Vivo
Determining the concentration of FtsZ is not straightforward. UV absorption is usually the simplest and most
accurate method [Brennan and Hardeman, 1993; Gill and
Von Hippel, 1989], but FtsZ has no tryptophan and almost
no tyrosine, and therefore has negligible absorption at
278 nm. Colorimetric assays can give widely different
color intensity for different proteins and need to be
standardized. We therefore used quantitative amino acid
analysis to determine the concentration of samples of
EcFtsZ, TmFtsZ, and BSA. (As an incidental result, we
independently determined the concentration of BSA by
UV absorption and confirmed that the classic extinction
coefficient, 0.667 [Foster and Sterman, 1956], is correct,
rather than the 7% lower value calculated from the amino
acid sequence [Brennan and Hardeman, 1993; Gill and
Von Hippel, 1989].) We then compared the three proteins
in the BCA assay and found that the ratio of color was
0.75 for EcFtsZ/BSA, and 0.67 for TmFtsZ/BSA. Therefore a substantial correction factor of 1.33 or 1.5 needs to
be applied when these proteins are measured in the BCA
assay, using BSA as a secondary standard. We also
calibrated the Bradford assay, in which the ratio of color
was 0.82 for EcFtsZ/BSA, and 0.70 for TmFtsZ/BSA. In
our experience, the BCA assay gives much more consistent results.
Two previous studies have reported the concentration of FtsZ in E. coli to be 20,000 or 5,000 molecules per
cell [Dai and Lutkenhaus, 1992; Pla et al., 1991], but the
methodologies were not well described. We therefore
determined the concentration of FtsZ in log phase BL21
E. coli by quantitative Western blotting, using the calibrated FtsZ sample described above. We averaged 12
quantitative blots over two separate experiments and two
different concentrations of cell lysate. The average value
was 14,800 molecules of FtsZ per cell, with a standard
deviation of 1,450. We conclude that there are approximately 15,000 molecules of FtsZ in the average log phase
E. coli cell, plus or minus 10%.
To compare our in vitro assays with conditions in
the cell, it is important to know the concentration of FtsZ
in the bacterial cytoplasm. The average E. coli cell (strain
B/r A, doubling time of 25min) has a length of 3.6 µm
[Donachie et al., 1976] and a diameter of approximately
0.94 µm [Trueba and Woldringh, 1980]. These dimensions
correspond well with our light microscopy observations
(data not shown). Assuming that the bacterial cell is a
cylinder capped by two hemispheres, the average cell volume is calculated to be 2.28 µm3. Kubitschek et al [1986]
gives the volume of a newborn cell under similar conditions as 1.566 µm3. An average cell under log phase growth
would have a volume of (1.566 µm3)/ln2, or 2.26 µm3,
which corresponds to our calculated volume. If there are
15,000 molecules of FtsZ in this volume, the intracellular
concentration of FtsZ is 10.9 µM, or 440 µg/ml.
We noted that the pET expression system for
EcFtsZ gives one of the highest expression levels of any
protein we have seen, so we determined the concentration
of the expressed protein. After 120 min induction we
found 3.1 3 106 molecules of FtsZ per cell, and noted
also that the cells were 2.4 times as long as normal log
phase cells. This gives an intracellular concentration of
36 mg/ml, or ,80 times the average normal concentration of FtsZ.
Stoichiometric GTP Binding
and Temperature-Dependent Hydrolysis
We used our calibrated protein concentration to
obtain an accurate measurement of the stoichiometry of
GTP binding to the three FtsZs. Starting with a sample of
FtsZ in 1 mM GTP, we removed free nucleotide by gel
filtration, determined the protein concentration as described, and precipitated the protein with 4.2% perchloric
acid. We then determined the concentration of guanine
nucleotide by UV spectrophotometry. We obtained a
value of 0.99 moles of guanine nucleotide per mol of
EcFtsZ. We did not determine whether the nucleotide was
GTP or GDP. The measured stoichiometry near 1:1
demonstrates that the affinity of guanine nucleotide is
sufficiently high to remain bound during gel filtration.
Similar measurements showed 1.03 mol guanine nucleotide per mol AzFtsZ and 1.13 mole guanine nucleotide
per mol TmFtsZ. This is the first study to use calibrated
protein concentration to demonstrate a 1:1 stoichiometry
of guanine nucleotide binding to FtsZ and confirms that
the FtsZ proteins purified by our protocols are ,100%
active in nucleotide binding.
We then assayed the GTPase activity in different
buffer conditions. We found that the GTPase activity of
both TmFtsZ and AzFtsZ required Mg21 (data not shown),
as shown previously for EcFtsZ [RayChaudhuri and Park,
1992; de Boer et al., 1992; Mukherjee et al., 1993] and
B. subtilis FtsZ [Wang and Lutkenhaus, 1993]. We
measured the GTPase activity of EcFtsZ in two assembly
buffers, MEMK6.5 and PB7.7. EcFtsZ showed measurable hydrolysis at 0°C, and a substantial increase at 37°C
(Fig. 1a). The activity in PB7.7 was higher than that in
MEMK6.5, probably because of the higher potassium
concentration, which enhanced activity significantly (see
below for details). The GTPase of TmFtsZ showed a
GTPase and Assembly of FtsZ
FtsZ, or (2) GTP turnover, in moles GDP produced per
mole FtsZ per min.
Requirement for Sodium or Potassium
for GTPase Activity
Mukherjee et al. [1993] reported that 100 mM
potassium was required for the GTPase activity of
EcFtsZ, and 2.0 M KCl was required for GTPase of the
halophilic archaebacterium Haloferax volcanii [Wang
and Lutkenhaus, 1996]. We investigated how the GTPase
of EcFtsZ varied with the concentration of KCl (Fig. 2a).
Minimal activation was obtained at 25 mM KCl, and
maximal stimulation required 100 mM KCl. Increasing
the KCl to 200–400 mM gave no additional enhancement.
400 mM KCl actually produced a several min lag before
hydrolysis began. 50 mM KCl gave about half-maximal
stimulation of GTPase, and this was used for most subsequent experiments. The enhancement was specific for potassium, since the GTPase in 50 mM NaCl was the same
as in Tris alone (Fig. 2b). This finding is consistent with
the previous study of EcFtsZ [Mukherjee et al., 1993].
However, we found that FtsZ from Azotobacter and
Thermotoga do not have a specific requirement for
potassium. As shown in Figure 2c,d, a monovalent cation
is required, but sodium is almost as effective as potassium
for AzFtsZ and is slightly more effective for TmFtsZ.
LiCl was also able to stimulate TmFtsZ GTPase activity
(data not shown).
GTPase Activity Depends on Concentration
of Protein and GTP
Fig. 1. GTPase activity of EcFtsZ and TmFtsZ at different temperatures. a: EcFtsZ (1.9 mg/ml) was tested at 0°C for 15 min; the
temperature was then shifted to 37°C. GTPase was tested in two
buffers. b: TmFtsZ was 0.8 mg/ml in TKM7.5. The plateau is reached
when all the GTP is hydrolyzed. Specific activity was expressed as
mols GTP hydrolyzed per mol FtsZ at each time point.
strong dependence on temperature (Fig. 1b). There was
very little activity at 37°C and a modest activity at 50°C.
At both 37°C and 50°C, there was a small lag in GTPase,
indicated by the acceleration of activity at later times.
GTPase was markedly increased and the lag was not
detectable at 65°C or higher.
Throughout this paper, we express GTP hydrolysis
as (1) specific activity, in moles GDP produced per mole
Our initial studies showed that the GTPase activity
was constant over a 5-fold concentration range, from 200
µg/ml to .1 mg/ml, for each of the three FtsZs. However,
previous studies have found that the GTPase activity of
FtsZ depends on the protein concentration, implying that
self-association may play a role in the hydrolysis reaction
[de Boer et al., 1992; Mukherjee et al., 1993; Wang and
Lutkenhaus, 1993]. In most cases, the major effect of
concentration was a steep loss of GTPase activity at FtsZ
concentrations of ,200 µg/ml, so we investigated this
range in detail.
Time courses of GTP hydrolysis at different FtsZ
concentrations are shown in Figure 3a, and the turnover
of GTP per FtsZ molecule per min in Figure 3b. Turnover
was independent of protein concentration above 300
µg/ml for Ec-, Az-, and TmFtsZ, and for all three proteins
the GTP turnover dropped below 200 µg/ml. This is
similar to BsFtsZ, which showed a similar plateau above
200 µg/ml, and a steep drop to zero activity at 100 µg/ml
[Wang and Lutkenhaus, 1993]. There were two different
behaviors at lower concentrations. For Ec- and AzFtsZ
there was a lag of 5–20 min before GTP hydrolysis began,
confirming two previous reports for EcFtsZ [de Boer et
Lu et al.
Fig. 2. Effect of monovalent cations on the GTPase activity of Ec-,
Az-, and TmFtsZ. Buffers all contained 70 mM Tris-HCl, pH 7.5,
5 mM magnesium acetate, 0.5 mM GTP, and either no additional salt,
or KCl or NaCl as indicated. a: Reaction containing 300 µg/ml EcFtsZ
and increasing amounts of KCl as indicated. b–d: The reactions
contained 50 mM KCl or NaCl as indicated (Tris indicates no
additional monovalent cation), plus (b) 650 µg/ml EcFtsZ; (c) 380
µg/ml AzFtsZ; (d) 800 µg/ml TmFtsZ. EcFtsZ and AzFtsZ were at
assayed 37°C, while TmFtsZ was at 65°C. The y-axis plots mols GTP
hydrolyzed per mol FtsZ at each time point. Each reaction reached a
plateau when all the GTP had been hydrolyzed. GTPase activity is
indicated by the initial slope, not the height of the curve at plateau.
Note that the time scale varies in the different panels.
al., 1992; Mukherjee et al., 1993]. For TmFtsZ there was
no lag, similar to reports for BsFtsZ [Wang and Lutkenhaus, 1993] and H. volcanii FtsZ [Wang and Lutkenhaus,
1996]. The lag suggests a nucleation phenomenon to
generate the active polymers, but it is curious that it is
observed in FtsZ from some species but not from others.
Our observation in a single experiment of a lag in two
species, and not in another, supports the interpretation
that these are real species differences and not due to
different protocols in different laboratories. We also
extend to three new species the observation that GTP
turnover per FtsZ is constant above 300 µg/ml, and drops
at lower concentrations, suggesting that this is a consistent feature of FtsZ from all bacteria.
The experiments depicted in Figure 3 were done at
GTP concentrations of 0.15–0.5 mM GTP, the lower
concentrations being used for increased accuracy at the
lower protein concentrations. Several points were done in
duplicate at both 0.15 and 0.5 mM GTP, and there was no
significant difference in the GTPase. Figure 4 confirms
GTPase and Assembly of FtsZ
that there is virtually no difference between 0.15 and 0.5
mM GTP. However, we found surprisingly large increases in GTPase when the GTP concentration was
increased to 1–5 mM (Fig. 4). This experiment used 0.3
mg/ml EcFtsZ, which was just above the plateau of the
curves in Figure 3, and either 50 or 350 mM KCl. There
were two effects of increasing GTP, and these were
similar at both KCl concentrations. First, there was no lag
at 0.2–0.5 mM GTP, but a 5- to 10-min lag appeared at
1–5 mM GTP. This is particularly evident in the reduced
hydrolysis at 5 min at the higher GTP concentrations.
Second, the rate of hydrolysis following the lag increased
markedly at higher GTP concentration. At each GTP
concentration the GTPase was about 2–3 times higher at
350 than at 50 mM KCl, similar to the results in Figure 2.
The maximum GTP hydrolysis obtained in our experiments was 26 mols GTP hydrolyzed per min per mol
EcFtsZ (at 350 mM KCl and 5 mM GTP). Figure 2
suggests that further increase in KCl would have no
effect, but Figure 4 suggests that hydrolysis rate would be
increased still further at higher GTP. Note that the GTPase
seems to be directly proportional to GTP over the concentration range 0.5–5 mM, but the curves do not extrapolate
to zero. At both 50 and 350 mM KCl, the GTP turnover
extrapolates to 2 min21 at low GTP concentration.
We considered the possibility that the enhanced
GTPase at 1–5 mM GTP could be a nonspecific effect of
nucleoside triphosphate. We tested the GTPase activity of
EcFtsZ in 0.5 mM GTP, with and without 4.5 mM ATP.
The GTPase activity was not affected by the 4.5 mM ATP.
We tested whether GDP, the product of GTPase,
could inhibit the GTPase activity of the FtsZ proteins. In
0.15 mM GTP and 300–500 µg/ml EcFtsZ, the addition of
an equal amount of GDP inhibited the GTPase activity
somewhat, and addition of a 5-fold excess of GDP
inhibited to 42% of full activity. AzFtsZ was more
sensitive to GDP than the others, and showed 68% (with
0.15 mM GDP) and 20% (with 0.75 mM GDP) of full
activity. TmFtsZ gave 78% activity in 5-fold excess GDP.
All Three FtsZs Assembled into Protofilament
Sheets or Bundles
To investigate further the oligomeric state of FtsZ
under nonassembly conditions, we sedimented protein
through glycerol gradients in 0.2 M ammonium bicarbonate. There is no GTPase activity in this buffer, and it does
not support assembly with DEAE dextran. Since TmFtsZ
shows neither GTPase nor assembly below 50°C, we
particularly expected this protein to be monomeric at
room temperature. Both Ec- and TmFtsZ sedimented near
the position of BSA, consistent with a monomer or dimer.
However, rotary shadowed electron micrographs showed
a substantial number of short straight protofilaments,
mixed with small globular particles that appeared to be
monomers (Fig. 5a,b). This association was most striking
for TmFtsZ, where the filaments were 5–10 subunits long.
This association must have occurred after the centrifugation because these longer filaments would have sedimented well ahead of BSA. Probably the association was
inhibited by the hydrostatic pressure in the sedimenting
gradient; a similar pressure inhibition was observed for
tubulin ring formation [Erickson, 1974]. Remarkably, the
assembly of TmFtsZ protofilaments in our ammonium
bicarbonate/glycerol buffer was essentially identical regardless of the addition of GTP, GDP, Mg or EDTA.
These results show that FtsZ from species as diverse as
E. coli and Thermotoga have a tendency to assemble into
straight protofilaments, even under conditions that block
GTPase and larger assembly.
We examined EcFtsZ by negative stain electron
microscopy in buffers promoting active GTPase and
found long, straight protofilaments in all three buffers
(Fig. 5c). Some protofilaments appeared to be single, and
others were paired. This assembly required GTP, but not
DEAE dextran. Protofilaments were formed at 0°C and
persisted during the first few min at 37°C. They disappeared after longer incubation at 37°C, probably when the
GTP was largely hydrolyzed. These long protofilaments
appear to be the same as those seen by Mukherjee and
Lutkenhaus by rotary shadowing [1994], and Wang et al.
[1997] by negative stain, and seem to be longer versions
of the short filaments in Figure 5a,b. These protofilaments
are good candidates for the oligomeric form required for
GTPase activity (see under Discussion).
Assembly of EcFtsZ protofilaments into larger
sheets could be obtained after addition of the polycation
DEAE dextran (Fig. 5d). The structure of these sheets and
their similarity to the microtubule wall has been described previously [Erickson et al., 1996]. Under some
conditions, EcFtsZ plus DEAE dextran produces small
tubes, similar to those described by Mukherjee and
Lutkenhaus [Mukherjee and Lutkenhaus, 1994]. The
structure of these tubes will be described elsewhere.
AzFtsZ in MEMK6.5 rapidly formed a white precipitate upon addition of Mg at 0°C. A similar but slower
precipitation occurred in PB7.7 after addition of Mg. This
precipitation required GTP, and therefore probably represents assembly into meaningful polymers. However, we
were not able to visualize these polymers by electron
microscopy, perhaps because they are too large. When
DEAE dextran was added before the Mg, the visible
turbidity was reduced and electron microscopy showed
loose protofilament bundles, with a less regular lattice
structure than that of EcFtsZ (Fig. 5e). We have yet not
found optimal conditions for assembling AzFtsZ.
Assembly of TmFtsZ required Mg, GTP, and high
temperature, but remarkably did not require DEAE
dextran. TmFtsZ in MEMK/Na6.5 plus GTP formed
Lu et al.
Figure 3.
GTPase and Assembly of FtsZ
highly ordered protofilament sheets with a very clear
resolution of individual protofilaments (Fig. 5f). TmFtsZ
sheets were temperature sensitive, and disassembled
quickly when the temperature dropped even to 50°C.
Fixation by glutaraldehyde was essential to preserve the
assembled sheets before preparation of negatively stained
specimens. Following assembly and fixation, more than
90% of TmFtsZ could be pelleted by centrifugation at
20,000g, indicating that the assembly was highly efficient.
We tested the effect of changing the concentration
of DEAE dextran on assembly and GTPase of EcFtsZ.
Figure 6a shows that in MEMK6.5 buffer 87% of the FtsZ
could be assembled and pelleted by centrifugation with a
ratio of 0.6 mg DEAE dextran/FtsZ. The fraction assembled dropped at both lower and higher DEAE dextran, very similar to the assembly of tubulin with DEAE
dextran [Erickson and Voter, 1976]. Electron microscopy
confirmed that polymers were similar in size at all DEAE
dextran concentrations, but much more numerous at 0.6
mg/ml. A reasonable interpretation is that DEAE dextran
is stabilizing the polymer by binding electrostatically to
and bridging two or more subunits. At lower concentrations of DEAE dextran, assembly is limited by the
stoichiometry: 0.06 mg/ml DEAE dextran assembled 0.5
mg FtsZ, compared with 0.42 mg tubulin [Erickson and
Voter, 1976]. In excess DEAE dextran, each DEAE
dextran molecule may bind a single FtsZ, rather than
bridging two or more and stabilizing the polymer. With
FtsZ, as with tubulin, the curves were similar at different
protein concentrations, i.e., it was the ratio of DEAE
dextran to protein that determined the polymerization.
Figure 6b shows that GTPase was inhibited by the
DEAE dextran-induced polymerization. A complication
in this experiment is that the polymer form changed. At
0.06 and 6 mg/ml, the polymers were protofilament
sheets, but at the peak assembly, 0.6 mg/ml DEAE
dextran, the polymers were tubes. It is likely that the tube
polymers are especially inhibited in GTPase, but the
protofilament sheets at 0.06 mg/ml are also inhibited
Fig. 3. Effect of protein concentration on GTPase activity in TKM7.5
buffer. a: Time course of GTP hydrolysis. Note again that the GTPase
activity is indicated by the initial slope of the curves, not by the height
of the plateau (which represents the exhaustion of all GTP). The
numbers to the right of each curve indicate the protein concentration in
µg/ml. The time scale on the x-axis is different for TmFtsZ. b: GTP
turnover (moles GTP hydrolyzed per mole of FtsZ per min) in the
interval of maximum hydrolysis. At the higher protein concentrations,
this was starting from the 0 time point, but at the lower protein
concentrations the turnover rate was determined after the lag (points in
parentheses). The GTP concentration was 0.15–0.5 mM, and several
duplicate experiments indicated no significant difference due to GTP
concentration in this range. EcFtsZ and AzFtsZ were assayed at 37°C,
TmFtsZ at 70°C. The straight lines fit the points, as well as more
complex curves.
relative to no DEAE dextran. It is important to remember
that FtsZ assembles into single or double protofilaments,
which appear to be the oligomer required for the high
GTPase, in the absence of DEAE dextran. These individual protofilaments were not pelleted in the centrifugation conditions used. Figure 6b indicates that further
assembly of these protofilaments into large sheets or tube
polymers substantially inhibits their ability to mediate
GTP hydrolysis.
Table I summarizes the GTPase activities of FtsZ
determined here and in several independent studies.
These hydrolysis rates, 2–10 GTP hydrolyzed per FtsZ
per min, are surprisingly high. Most proteins that hydrolyze GTP have a very low intrinsic GTPase, which is
accelerated by interaction with other proteins. In a
general discussion of the GTPase superfamily, Bourne et
al. [1991] comment: ‘‘In many (but not all) GTPases the
intrinsic rate constants of GDP release (kdiss.GDP) and GTP
hydrolysis (kcat.GTP) are quite low (,0.03 min−1).
These are increased by either of two classes of regulatory
proteins: the guanine nucleotide release proteins (GNRPs),
which catalyze release of bound GDP, promoting its
replacement by GTP, and the GTPase activating proteins
(GAPs), which speed up GTP hydrolysis.’’
GAPs can accelerate GTP hydrolysis up to 5 orders
of magnitude when they bind to their cognate G protein.
The mechanism has been revealed most clearly in the
recent x-ray structures of Ras●RasGAP [Scheffzek et al.,
1997], and Rho●RhoGAP [Rittinger et al., 1997], which
show that the GTP molecule is in the interface between
the G-protein and the GAP, and makes contact with both
proteins. Importantly, most catalytic residues are provided by Ras or Rho, but in both cases one catalytic
residue is provided by the GAP. The GAP may also
stabilize a conformation of the G-protein favoring the
transition state.
Tubulin has a very low GTPase activity under most
conditions that prevent its assembly into microtubules,
but during microtubule assembly each tubulin subunit
hydrolyzes one GTP shortly after it enters the microtubule lattice. We have postulated that tubulin acts as its
own GAP, stimulating hydrolysis only when two subunits
encounter each other for productive assembly [Erickson
and O’Brien, 1992]. The GTPase of dynamin has some
similarity to that of tubulin. The basal GTP turnover is
much higher, about 2 min21 but, like tubulin, it is
significantly enhanced to 10 min21 when assembled into
polymers [Warnock et al., 1996]. Cooperative assembly
onto microtubules or lipid vesicles can increase the
activity to 100 min21 [Tuma et al., 1993]. Thus, the
Lu et al.
Fig. 4. Effect of GTP concentration on GTPase activity of EcFtsZ. The buffer was as in Fig. 3, except for
350 mM KCl (bottom). EcFtsZ was 0.3 mg/ml in all cases. Left, top and bottom, GTP hydrolysis as a
function of time at different GTP concentrations, indicated in mM on the right of each curve. Right,
maximum GTP turnover, measured after the lag when one exists.
GTPase activity of dynamin is expressed primarily when
it is assembled into its characteristic polymer, which in
this case is a curved protofilament forming rings or stacks
of rings [Hinshaw and Schmid, 1995].
An important characteristic of tubulin GTPase is
that subunits in the interior of the microtubule cannot
exchange nucleotide, so hydrolysis only occurs when
new subunits are added to the ends. Since GTP hydrolysis
is coupled to assembly, the GTPase is proportional to the
concentration of soluble tubulin and the concentration of
microtubule ends. GTP hydrolysis can reach 1–3 min–1
per mol of total tubulin during the initial phase of
microtubule assembly, but slows to 0.06 min–1 at steady
state [O’Brien et al., 1987]. Obviously, steady-state
hydrolysis requires a continuous association of subunits
onto some microtubules, which must be balanced by an
equal dissociation of subunits from other microtubules.
This process is known as microtubule dynamic instability
[Erickson and O’Brien, 1992]. Continuous assembly and
disassembly of dynamin has also been implicated in its
GTPase [Warnock et al., 1996], but the mechanism is
most clearly established for microtubules.
GTPase and Assembly of FtsZ
Fig. 5. a,b: Electron microscopy of FtsZ after glycerol gradient
sedimentation. Rotary shadowed specimens were prepared at room
temperature from gradient fractions in 0.2 M ammonium bicarbonate,
30% glycerol. EcFtsZ (a) shows mostly small globular subunits that we
interpret to be FtsZ monomers (arrow), and some dimers and trimers.
TmFtsZ (b) shows some monomers (arrow), but mostly short straight
protofilaments several subunits in length. c: Single (arrow) and paired
(arrowhead) protofilaments imaged by negative stain, from samples of
EcFtsZ under conditions of high GTPase. d–f: Protofilament sheets and
bundles assembled by EcFtsZ, AzFtsZ, and TmFtsZ. d: EcFtsZ (1
mg/ml) plus 0.6 mg/ml DEAE dextran in MEMK6.5 at 37°C assembles
into protofilament sheets. Similar sheets were assembled in TKM7.5
and PB7.7. e: AzFtsZ was assembled with 1 mg/ml protein plus 0.6
mg/ml DEAE dextran at 37°C in PB7.7. It forms bundles of protofilaments in a looser array than EcFtsZ sheets. f: TmFtsZ assembled at 1
mg/ml protein and no DEAE dextran at 70°C in MEMK6.5. This
preparation was fixed by adding glutaraldehyde to 1%, and a negatively
stained specimen was prepared after 5 min at 70°C.
Lu et al.
Fig. 6. Assembly and GTPase assay of FtsZ in MEMK6.5 containing 1
mg/ml FtsZ, 1 mM GTP, and DEAE dextran as indicated. a: Assembly
of FtsZ at 37°C for 5 min was assayed by centrifugation at 20,000g,
4°C. The pelleted protein was dissolved in 0.1 M NaOH and its
concentration determined by the BCA protein assay. The assembly is
presented as percentage of FtsZ pelleted. b: GTPase assay of FtsZ
under the same conditions. Reactions were assayed every 3 min, and
the GTPase activity expressed as turnover of GTP per FtsZ per min. SD
were calculated from three (a) or two (b) independent experiments.
GTP Hydrolysis Is Coupled to FtsZ Polymerization
The sigmoidal dependence of GTPase on FtsZ
protein concentration was first observed for FtsZ from
B. subtilis [Wang and Lutkenhaus, 1993]. At ,80 µg/ml,
the GTPase was essentially zero, but at slightly higher
concentrations the specific activity increased very steeply.
At 80–150 µg/ml, the GTPase increased from 1.3 to 9.3
min21; then, at 150 µg/ml, the specific activity abruptly
plateaued and remained constant at higher concentrations. Wang and Lutkenhaus [1993] concluded that ‘‘the
B. subtilis FtsZ must oligomerize in order to express
GTPase activity.’’ FtsZ from Haloferax volcanii showed
similar behavior at low concentrations: virtually no
GTPase at 100 µg/ml, increasing steeply from 0.24 to 1.3
min21 as the concentration increased from 200 to 400
µg/ml [Wang and Lutkenhaus, 1996] (higher concentrations were not explored, so a plateau was not determined).
We now report that Ec-, Az-, and TmFtsZ show a
concentration dependence of GTPase very similar to
BsFtsZ: a low GTPase at ,50 µg/ml protein concentration, increasing sharply from 50 to 300 µg/ml, and a
plateau, where the specific GTPase is independent of
protein concentration, of .300 µg/ml. The most reasonable explanation is the one suggested previously, that the
GTPase activity is expressed only when FtsZ forms an
oligomer. The sigmoid shape and steepness of the activation with concentration strongly suggest a nucleated
polymerization, in which the oligomer is higher order
than a dimer. The abrupt plateau in specific activity at
.200–400 µg/ml implies that above this concentration
virtually all additional protein associates into oligomers
with the same GTPase activity. It is interesting that the
average concentration of FtsZ in E. coli was found to be
440 µg/ml, somewhat above this plateau value.
The best candidate for the active oligomers are the
single and paired protofilaments seen by negative stain
electron microscopy (Fig. 5c). These protofilaments were
abundant at protein concentrations of .300 µg/ml, and
rare at ,50 µg/ml, consistent with the concentration
range for formation of the active oligomeric species.
The sigmoidal curve implies a multi-subunit nucleated assembly, and this is also suggested by the lag in
GTPase seen at low protein concentration, at high GTP,
and at 400 mM KCl. However, nucleated assembly would
not be expected for single protofilaments. Addition of a
subunit to the end of a protofilament should involve the
same reaction as formation of a dimer, so assembly of a
single protofilament should be an isodesmic reaction with
no nucleation phase. It is therefore likely that the active
polymers are double protofilaments or larger parallel
arrays. Indeed, many double protofilaments are seen in
the micrographs, and some images that appear to show
single protofilaments may be edge views of double
filaments. The electron microscopy so far is suggestive
but preliminary, and more will be needed to specify the
structure of the active polymers.
The Question of Nucleotide Exchange and GTPase
Activity of Larger Polymers
Continuous GTP hydrolysis requires that GDP be
exchanged for GTP following hydrolysis, to initiate a
second round. Since hydrolysis appears to be associated
with polymers, there are two possible mechanisms for
this exchange. One possibility is internal exchange,
where the nucleotide would be exchanged from every
subunit in the polymer. With microtubules this does not
occur; the nucleotide is trapped in the interface between
GTPase and Assembly of FtsZ
Spec. act.
Temp. (°C)
Protein concn. (mg/ml)
KCl (mM)
GTP (mM)
50 mM Tris-HCl, pH 7.5, 50 mM KCl, 5 mM MgAC2 , 0.15–0.5 mM GTP; B, 50 mM Hepes, pH 7.2, 100 mM KCl, 10 mM MgCl2 , 0.5 mM
GTP; C, 40 mM Tris-acetate, pH 7.0, 200 mM KAC, 15 mM MgAC2 , 5 mM GTP, 2 mM EDTA, 1 mM DTT, 0.5% Triton X-100; D, 50 mM
Tris-HCl, pH 7.5, 50 mM KCl, 2.5 mM MgCl2 , 0.5 mM GTP, 5% glycerol; E, 50 mM Hepes, pH 7.2, 2 M KCl, 10 mM MgCl2 , 0.5 mM GTP.
TmFtsZ, Thermotoga maritima FtsZ; EcFtsZ: Escherichia coli FtsZ; AzFtsZ, Azotobacter vinelandii FtsZ; BsFtsZ: Bacillus subtilis FtsZ; HvFtsZ,
Haloferax volcanii FtsZ.
bFtsZ was preincubated with 5 mM GTP at 37°C for 80 min.
ca, this work; b, Fig. 4 of Mukherjee [1993]; c, Fig. 3a of de Boer et al. [1992]; d, Fig. 3b of de Boer et al. [1992]; e, Fig. 2e of Ray Chaudhuri
[1992]; f, Fig. 7A of Wang et al. [1993]; g, Fig. 2B of Wang et al. [1996].
subunits, so that internal subunits cannot exchange nucleotide. The other possibility is exchange on free subunits
accompanied by continuous dissociation and reassociation. With microtubules, nucleotide hydrolysis occurs
rapidly after a subunit associates onto the end of a
microtubule, and the subunit retains its GDP as long as it
remains in the microtubule. During the shortening phase
of dynamic instability subunits are released, and once free
in solution they can exchange GDP for GTP and be
charged for another round of assembly. It is unclear how
this scenario might apply to FtsZ, but the similarity of
atomic structures [Löwe and Amos, 1998; Nogales et al.,
1998] strongly suggests that the nucleotide will be
nonexchangeable in the straight protofilament.
We observed in Fig. 6 that assembly into larger
sheets of protofilaments stabilized by DEAE dextran
substantially reduced the GTPase activity. Yu and Margolin [1997] reported recently that 10 mM calcium induced
assembly of larger polymers; we have repeated these
results (unpublished work) and found that calcium induces assembly of protofilament sheets and tubes that are
essentially identical to those induced by DEAE dextran.
Yu and Margolin also reported reduction in GTPase at the
calcium concentrations giving maximum assembly, similar to our results with DEAE dextran. Apparently the GTP
hydrolysis is stimulated primarily by small polymers, the
protofilaments shown in Figure 5c, and inhibited when
these protofilaments associate into larger polymers. A
likely explanation is that dissociation and turnover of
subunits is reduced in the larger polymers.
A final word of caution is needed concerning the
high rate of GTP hydrolysis observed in vitro. Hydrolysis
of 2–10 molecules of GTP per min by the 15,000
molecules of FtsZ in a cell would seem a large metabolic
burden. Little is known about the dynamics of FtsZ
assembly in the cell, so perhaps it is assembled once and
maintained in a polymer form that blocks GTP exchange
and further hydrolysis. A different enigma is posed by the
mutant FtsZ84. This temperature sensitive mutant is
defective for cell division at 42°C but can function fully
at 30°C. However, its GTPase in vitro is less than
one-tenth that of wild-type FtsZ over the entire range of
30–44°C [RayChaudhuri and Park, 1992; de Boer et al.,
1992]. This finding suggests that the very high level of
GTP hydrolysis observed in vitro may not be biologically
important. The lower hydrolysis rates achieved by FtsZ84,
or by FtsZ assembled with DEAE dextran may be more
relevant to the in vivo function.
We thank Dr. David Bramhill, Merck, for providing
the pET plasmid for expression of EcFtsZ; Dr. Michael
Adams, University of Georgia, for Thermotoga DNA; Dr.
Lu et al.
Paul E. Bishop, North Carolina State University, for
Azotobacter DNA; Dr. Limin Zhang, Virginia Polytechnic Institute, for the Azotobacter cosmid library.
Addinall, S.G., Bi, E.F., and Lutkenhaus, J. (1996): FtsZ ring formation
in fts mutants. J. Bacteriol. 178:3877–3884.
Beall, B., Lowe, M., and Lutkenhaus, J. (1988): Cloning and characterization of Bacillus subtilis homologues of Escherichia coli cell
division genes ftsA and ftsZ. J. Bacteriol. 170:4855–4864.
Bourne, H.R., Sanders, D.A., and McCormick, F. (1991): The GTPase
superfamily: Conserved structure and molecular mechanism.
Nature 349:117–127.
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