Download A Dictyostelium Mutant with Reduced Lysozyme Levels

THE JOURNAL OF BIOLOGICAL CHEMISTRY
© 2005 by The American Society for Biochemistry and Molecular Biology, Inc.
Vol. 280, No. 11, Issue of March 18, pp. 10435–10443, 2005
Printed in U.S.A.
A Dictyostelium Mutant with Reduced Lysozyme Levels
Compensates by Increased Phagocytic Activity*
Received for publication, October 7, 2004, and in revised form, December 28, 2004
Published, JBC Papers in Press, January 7, 2005, DOI 10.1074/jbc.M411445200
Iris Müller‡, Ninon Šubert§, Heike Otto‡, Rosa Herbst¶储, Harald Rühling‡, Markus Maniak‡**,
and Matthias Leippe‡‡
From the ‡Department of Cell Biology, Kassel University, Heinrich-Plett-Strasse 40, 34132 Kassel, §Bernhard Nocht
Institute for Tropical Medicine, Bernhard-Nocht-Strasse 74, 20359 Hamburg, ¶Department of Special Zoology, Ruhr
University of Bochum, Universitätsstrasse 150, 44780 Bochum, and ‡‡Zoological Institute of the University of Kiel,
Olshausenstrasse 40, 24098 Kiel, Germany
Lysozymes are bacteria-degrading enzymes and play a
major role in the immune defense of animals. In freeliving protozoa, lysozyme-like proteins are involved in
the digestion of phagocytosed bacteria. Here, we purified a protein with lysozyme activity from Dictyostelium
amoebae, which constitutes the founding member, a
novel class of lysozymes. By tagging the protein with
green fluorescent protein or the Myc epitope, a new type
of lysozyme-containing vesicle was identified that was
devoid of other known lysosomal enzymes. The most
highly expressed isoform, encoded by the alyA gene, was
knocked out by homologous recombination. The mutant
cells had greatly reduced enzymatic activity and grew
inefficiently when bacteria were the sole food source.
Over time the mutant gained the ability to internalize
bacteria more efficiently, so that the defect in digestion
was compensated by increased uptake of food particles.
The lysosome is the most potent degradative organelle
within the eukaryotic cell. It contains hydrolytic enzymes that
fulfill essential functions. In humans many mutations affecting
its constituents lead to the diseased state, often with dramatic
consequences. Mutations of this sort fall into two classes. The
first class affects proteins involved in trafficking to and from
the lysosome and/or regulating its integrity by modulating
fusion and fission events. Among these proteins are Rabs,
proteins belonging to a family that act as molecular timers (1)
controlling virtually every trafficking step in the cell (2) and
the LYST proteins, which when mutated cause enlarged lysosomes manifested as Chediak-Higashi Syndrome (3). The second class of mutations affects lysosomal enzymes with metabolic roles. These enzymes degrade macromolecules that the
cell wants to dispose of, producing small metabolites that are
useful building blocks or provide energy if completely degraded
in the cytosol and mitochondria. Tragically, many of the lysosomal enzymes are exoenzymes, i.e. they digest macromolecules from their ends. If one of these enzymes is lacking, degradation proceeds up to the residue that cannot be removed,
and as a consequence, all steps that ensue will be blocked. This
* This work was supported by the Deutsche Forschungsgemeinschaft
Grants MA 1439/4-1 and LE 1075/2-3. The costs of publication of this
article were defrayed in part by the payment of page charges. This
article must therefore be hereby marked “advertisement” in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
储 Recipient of a Lise Meitner fellowship from Nordrhein-Westfalen.
** To whom correspondence should be addressed: Abt. Zellbiologie,
Universität Kassel, Heinrich-Plett-Str. 40, 34132 Kassel, Germany.
Tel.: 49-561-804-4798; Fax: 49-561-804-4592; E-mail: maniak@
uni-kassel.de.
This paper is available on line at http://www.jbc.org
leads to the accumulation of partially degraded intermediates
within the lysosomes culminating either in the formation of
toxic compounds or blowing up the volume of the lysosome until
it sterically interferes with vital cellular activities. Depending
on the cell type that is most seriously harmed, be it neurons,
muscle, or leukocytes, patients will suffer from mental retardation, cardiac failure, or immunodeficiency (4, 5).
In contrast, free-living amoebae are unlikely to develop lysosomal storage diseases because their endocytic pathway differs from that of mammalian cells. In mammalian cells, the
lysosome is a dead-end of vesicular trafficking. In most cell
types, it accumulates non-degradable material and retains it
for the lifetime of the cell and organism. In amoebae, endocytosed cargo transits through the cell. Any material that cannot
be degraded because specific enzymes are lacking is released
from the cell by exocytosis after an hour. Despite these differences, amoebae such as Dictyostelium can serve as bona fide
model systems to study lysosomal function in vivo. Interference
with Rabs (6, 7) and LYST proteins (8, 9) produces phenotypes
in Dictyostelium that are comparable with those observed in
mammalian cells, indicating that many lysosomal constituents
are functionally conserved in evolution. On the other hand, a
deficiency of cathepsin D leads to profound defects in mice (10)
and sheep (11) but is tolerated well in Dictyostelium (12).
More recently mice have been generated that specifically
lack the M isoform of lysozyme, which constitutes the most
prominent bacteriolytic activity in the airways (13). Although
homozygous mutants increase the level of expression of the
lysozyme P isoform to compensate for their deficiency, they
remain much more sensitive to infections of the lung (14).
Lysozymes are not only found in animals but are also present
in numerous phylogenetically diverse organisms such as
plants, fungi, bacteria, and bacteriophages (15). Several different classes of theses enzymes have been described revealing
that structurally diverse proteins fulfill the function of degrading the peptidoglycan of bacteria by splitting a 1,4-linkage
between N-acetylmuramic acid and N-acetylglucosamine (16).
Despite the fact that Dictyostelium has been regarded as a good
model for elucidating molecular mechanisms underlying phagocytosis of cells from higher organisms, e.g. mammalian defensive cells, nothing is known about the molecular armament
that it uses to efficiently eliminate the enormous number of
bacteria phagocytosed for nutrition. We have isolated a protein
with lysozyme activity from Dictyostelium amoebae. The protein is the first antimicrobial polypeptide characterized in this
protozoan organism at the molecular level, and it appears to be
a member of a previously unrecognized lysozyme family. The
enzyme is stored in a unique organelle, and disruption of its
gene had a dramatic effect on the phenotype.
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Compensation of Lysozyme Deficiency
EXPERIMENTAL PROCEDURES
Purification of Lysozyme—The AX2 strain of Dictyostelium discoideum was cultured at 22 °C axenically (17). Amoebae in late-logarithmic phase were harvested, sedimented at 320 ⫻ g at 4 °C for 5 min, and
washed 2 times in Sörensen’s 17 mM sodium/potassium phosphate, pH
6.0. Freshly harvested and washed amoebae (2,5 ⫻ 109 cells) were lysed
by three cycles of freezing and thawing in 5 volumes of 10 mM sodium
acetate, pH 4.5, supplemented with complete proteinase inhibitor mixture (Roche Applied Science). The lysate was centrifuged at 150,000 ⫻
g at 4 °C for 1 h. Subsequent purification steps were carried out at
10 °C, and samples were kept on ice. The 150,000 ⫻ g supernatant was
applied to a chromatography column (Econo column 2.5 ⫻ 20 cm,
Bio-Rad) manually filled with a Bio-Gel matrix (Bio-Gel A-0.5m; BioRad). The column was equilibrated with 10 mM sodium acetate, pH 4.5.
Adsorbed protein was eluted by washing the column with the same
buffer (350 ml) and with 100 mM sodium acetate, pH 4.5 (200 ml).
Finally, the column was washed with 500 mM sodium acetate, pH 4.5
(200 ml). Fractions with lysozyme activity were pooled and loaded on a
Resource S cation exchange column (1 ml; Amersham Biosciences)
equilibrated with 10 mM sodium acetate, pH 4.5. Adsorbed protein was
eluted by first washing the column with the same buffer (5 ml), then by
use of a gradient from 0 to 300 mM NaCl (25 ml) and finally by a wash
with 1 M NaCl. Each fraction was tested for lysozyme activity.
Protein Analysis—Protein concentration was determined using the
microbicinchoninic acid reagent assay (Pierce) and hen egg lysozyme as
standard (18). Tricine1 SDS-PAGE was performed according to Schägger
and von Jagow (19). For N-terminal sequencing proteins were reduced
with dithiothreitol and alkylated with iodoacetamide, subjected to Tricine
SDS-PAGE and subsequent semidry electroblotting using a polyvinylidene difluoride membrane (Problott; Applied Biosystems), and analyzed
using a gas-phase protein sequencer (model 437A; Applied Biosystems).
For the detection of glycosylation proteins were analyzed after blotting
onto nitrocellulose membrane using a glycan detection assay according to
the manufacturer’s instructions (Roche Applied Science). For mass spectrometric analysis, samples were dialyzed against 5% acetic acid and
mixed with the same volume of saturated trans-sinapinic acid in acetonitrile, 0.1% trifluoroacetic acid as UV-absorbing matrix. About 10 ␮l of the
samples were spotted on a stainless steel probe tip and dried at room
temperature. Measurements were performed using a Bruker Relex
MALDI-TOF. Ions were formed by laser desorption at 337 nm using an N2
laser. Spectra were recorded with an acceleration voltage of 35 kV in the
linear mode. The mass spectrometer was calibrated with bovine serum
albumin and carbonic anhydrase from bovine erythrocytes (both from
Sigma) as external standards.
Enzyme Assays—Lysozyme activity was measured as described previously (20). In all assays hen egg lysozyme (Sigma) served as a control. To
monitor lysozyme activity during purification, column fractions were analyzed using the lysoplate technique (21). For quantitative measurements
of lysozyme activity, cell wall degradation of lyophilized Micrococcus luteus (Sigma) was determined turbidimetrically essentially according to
Shugar (22). Degradation of cell walls of Staphylococcus aureus (Sigma)
was tested in the same assay. To determine the effect of pH and ionic
strength on lysozyme activity, the turbimetric assay was adjusted according to Dobson et al. (23). Various buffers were prepared as specified by
Miller and Golder (24). They covered a pH range from 3.5 to 7.5, their ionic
strength varying between 0.005 and 0.2. The occurrence of reducing
N-acetyl amino sugars was tested as described by Reissig et al. (25).
Chitinolytic activity was assayed using chitin glycol (Sigma) as a substrate (26). Activity against viable bacteria was determined in a microdilution susceptibility assay as published previously (27).
Gene Disruption—The sequence encoding AlyA was amplified from
AX2 genomic DNA using primers bearing an EcoRI recognition sequence in front of the start codon (5⬘-CCGGAATTCAAAATGAGAATTGCTTTCTTTTTACTTGTTTTAGCCG-3⬘) and a BglII site after the
last codon specifying an amino acid (5⬘-GGAAGATCTTTCGAGATCAAAACCATTACAGGTCTTTTCAGC-3⬘), cleaved with the corresponding enzymes, and ligated into the same sites of pIC19H. After
deleting a SalI site in the polylinker of this plasmid, which is also
recognized by HincII, the lysozyme sequence was cut at its remaining
HincII site, and a blunted cassette conferring blasticidin resistance,
isolated via BamHI and HindIII digestion from plasmid pUCBsrDel1
The abbreviations used are: Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; Aly, amoeba lysozyme; mAb, monoclonal
antibody; GFP, green fluorescent protein; MALDI-TOF, matrix-assisted
laser desorption ionization time-of-flight; TRITC, tetramethylrhodamine isothiocyanate.
Bam (28), was inserted. The resulting vector was cleaved with SmaI
and NruI situated in the polylinker region to release the plasmid
backbone required for replication and selection in Escherichia coli.
The remaining linear 2.5-kilobase fragment containing 560 bp of
lysozyme homologous sequences upstream and 550 bp downstream of
the resistance cassette was transfected into Dictyostelium AX2 cells
by electroporation, and the resulting clones were isolated by limited
dilution into multiwell plates. To identify clones that carry a disrupted lysozyme gene, we performed PCR on preparations of genomic
DNA (29) using primers (5⬘-GTGGTGCTCTCTATTTCACAGC-3⬘ and
5⬘-CTTGTTCAACCCACATGGCTGG-3⬘) that flank 85 bp of the coding sequence containing two introns of different lengths which distinguish between the genes encoding lysozyme isoforms.
Expression of Lysozyme and Tagged Variants—To rescue possible
defects in a lysozyme mutant, EcoRI and HindIII enzymes were used to
cut out lysozyme from pIC19H, and the gene was inserted into the
expression vector pDEX-RH (30) cut with the same restriction enzymes.
To allow for detection of the transgene, a Myc epitope sequence cloned
in pIC20R (31) was cut out using SmaI and AccI enzymes, blunted by
filling in the two-base overhang, and ligated into the HincII site within
the lysozyme sequence of the pDEX-RH expression construct. To monitor the distribution of lysozyme, a GFP fusion was constructed. The
lysozyme coding sequence from the ATG up to the cleavage site of the
prodomain was amplified by PCR using a cDNA library in phage ␭
(courtesy of Peter Devreotes) as a template. The product was digested at
the EcoRI and BglII cleavage sites that flanked the PCR primers 5⬘-CCGGAATTCAAAATGAGAATTGCTTTCTTTTTACTTGTTTTAGCCG-3⬘ and 5⬘-GGAAGATCTTGTAACAGTTGCTGTGATAAGGAAATGATCAC-3⬘ and inserted into pDd-A15-gfp (32) as modified by (33), so that
GFP replaces the C-terminal prodomain of lysozyme.
Endocytosis and Cell Growth—The uptake of particles and fluid
phase was measured as described previously (34, 35). To determine the
capacity of cells to degrade bacteria in vivo, we used the assay established by Maselli et al. (36), except that we used E. coli expressing the
green fluorescent protein (37) to measure the decrease in intracellular
fluorescence after phagocytosis. To assay the growth properties of cells
on bacterial lawns, we used M. luteus, Klebsiella aerogenes, and E. coli
as substrates. These bacterial species were pre-grown on SM plates,
harvested in Sörensen’s phosphate buffer (as above), and mixed with
about 80 cells of the appropriate Dictyostelium strain, and plaque
diameter was measured after 5 days of growth on SM agar plates. In
this assay no dramatic growth differences were observed between
Gram-negative and Gram-positive bacteria.
Immunofluorescence and Antibodies—mAb 221-342-5 recognizes a
sulfated mannose determinant residing on many lysosomal enzymes
(38). mAb AD 7.5 binds to an N-acetylglucosamine 1-phosphate modification of another class of lysosomal enzymes (39). The antigen corresponding to mAb 130-80-2, an esterase gp70, accumulated in crystal
bodies (40). mAb 9E10 recognizes the Myc epitope (41). The anti-GFP
antibody (264-449-2) is available from Chemicon (Temecula, CA). Immunofluorescence experiments were performed as described before (34).
Monoclonal antibodies were detected using TRITC-coupled polyclonal
secondary antibodies purchased from Dianova (Hamburg, Germany).
Images were taken from single confocal planes using a Leica TCS-SP
laser-scanning microscope.
Data Base Searches and Accession Numbers—A computer-assisted
homology search was performed using the Internet BLAST and FASTA
searches in the nucleic acid data base of the National Center for Biotechnology Information (NCBI) and the Dictyostelium protein data base
(dictybase.org/db/cgi-bin/blast.pl), resulting in sequence information for
alyA (AAM08434), alyB (AAM08432), alyC (AAO051440), alyD
(DDB0217279 and DDB016810), DdLysC1, DdLysC2, and DdLysC3
(lysozymes of D. discoideum similar to c-type lysozymes; U66523,
DDB0189256, DDB0205537, respectively). Dictyostelium genes coding
for DdLys T41 and DdLysT42 (lysozymes of D. discoideum similar to T4
phage-type lysozymes; DDB0217501 and DDB0167824, respectively)
and DdLysEh1 and DdLysEh2 (lysozymes of D. discoideum similar to
Entamoeba histolytica; DDB0167552 and DDB0219864, respectively)
were also identified in the Dictyostelium genome. The amino acid sequences of lysozymes of diverse organisms used in the phylogenetic
analysis were retrieved from the Protein Entrez data base of NCBI and
listed below with their respective abbreviations in Fig. 7 (source organism; accession numbers): LysC (Gallus gallus c-type lysozyme;
2CDS_A), LysT4 (enterobacteria phage T4-type lysozyme; 1LYD), LysG
(goose-type lysozyme; LZGSG). The following lysozymes represent the
i-type lysozymes of invertebrates: Bathymodiolus azoricus (AAN16208),
Bathymodiolus thermophilus (AAN16209). (Caenorhabditis elegans 1–3
(AAC10179, AAC19181, AAA83197), Calytogena sp. 1 and 2
Compensation of Lysozyme Deficiency
FIG. 1. Purification of AlyA and activity determination at various conditions. A, a protein preparation with lysozyme activity
eluted isocratically from a Bio-Gel column was subjected to Resource S
cation exchange chromatography. Bound protein was eluted with an
increasing NaCl gradient (dotted line), and active fractions were pooled
as indicated by the bar. The inset shows the different steps of purification of AlyA on a silver-stained SDS gel. Lane 1, amoebic acid extract;
lane 2, Bio-Gel fraction; lane 3, Resource S fraction. Molecular masses
are indicated at the left. B, cell walls of M. luteus were incubated with
the purified protein at various pH and at ionic strength reaching from
0.005 to 0.2, and degradation as a measure of lysozyme activity was
determined photometrically.
(AAN16211, AAN16212), Chlamys islandica (CAB63451), Drosophila
melanogaster 1-3 (CAA21317, AAF57939, AAF57940), Hirudo medicinalis (AAA96144), Mytilus galloprovincialis (AAN16210), Mytilus
edulis (AAN16207), Tapes japonica (BAB3389), Asterias rubens
(AAR29291), E. histolytica EhLys 1 and 2 (Q27650, AAC67235).
Protein alignments were performed using the software ClustalX (42),
and phylogenetic relationships were analyzed using PAUP 4.0 (43).
Bootstrap analysis (1000 replicates) was added to test tree robustness.
RESULTS
Purification of a Novel Lysozyme and Identification of the
alyA Gene—We purified a protein with lysozyme activity from
axenically cultured Dictyostelium amoebae, exploiting its adsorption to a Bio-Gel matrix. The main part of active material
found in acid extracts of amoebae was only weakly retained by
the column. It was eluted as a single protein peak with the
starting buffer. Final purification was achieved by cation exchange chromatography and yielded homogeneous material
with a molecular mass of ⬃12 kDa as judged by SDS-PAGE
(Fig. 1A). The purified protein degraded M. luteus cell walls
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optimally at low ionic strength between pH values ranging
from 4.5 to 6 (Fig. 1B). A colorimetric test for detecting reducing ends of N-acetyl amino sugars within a preparation of
degraded cell walls was positive and comparable to hen egg
lysozyme. Because the protein did not exert chitinase activity
in the detection system used (data not shown), these results
together support a true N-acetylmuraminidase (lysozyme) activity of the protein. Purified lysozyme was relatively stable; at
pH 5.5, 10-min incubations at 50 °C did not reduce the activity,
but in contrast to hen egg white lysozyme, its activity decreased to half when the enzyme was incubated at 60 °C for 10
min. The enzyme displayed antibacterial activity against viable bacteria; the protein inhibited the growth of the Grampositives Bacillus subtilis and Bacillus megaterium at 2.5 ␮g/
ml. The growth of E. coli, representing Gram-negatives, was
not inhibited at 10 ␮g/ml. Neither viable S. aureus nor cell
walls of this bacterium were affected by the enzyme at this
concentration (data not shown).
The purified lysozyme was subjected to N-terminal sequencing, yielding a single amino acid sequence up to residue 25
(YSCPKPCYGNMCCSTSPNNQYYLTD). This sequence was
back-translated into nucleic acid sequence and used to search a
cDNA data base (44). A cDNA clone, SSF469, was retrieved, the
deduced amino acid sequence of which perfectly matched the
experimentally determined amino acid sequence of lysozyme.
Using this clone we could identify the corresponding gene on
chromosome 2 in the data base from the Dictyostelium genome
project (45) and named it alyA (for amoeba lysozyme ). The
information on genomic structure and amino acid sequence is
summarized in Fig. 2.
The N-terminal tyrosine residue of the mature enzyme is
preceded by a stretch of 19 mostly hydrophobic amino acid
residues that may serve as a signal peptide necessary for the
transport of the protein into the endoplasmic reticulum membrane. MALDI-TOF analysis of the purified AlyA protein
yielded a molecular mass of 12.652 kDa. This value confirms
the molecular mass estimated from SDS-PAGE and is in good
agreement with a mature peptide of 119 residues that would
have a calculated molecular mass of 12.633 kDa provided that
the 12 cysteine residues participate in disulfide bonding. No
glycosylation of the purified protein was experimentally evident as judged by a glycan detection assay. The only putative
N-glycosylation site in the primary translation product is at
Asn149, beyond the C terminus of the mature protein. Neither
the mature protein nor its precursor molecule reveal significant
similarity to any protein of other organisms reported so far, in
particular not to other bacteriolytic enzymes.
Phenotypic Instability of aly Knockouts—To analyze the in
vivo function of this novel protein, we disrupted the alyA gene.
Therefore, we electroporated Dictyostelium cells in the presence of a linear DNA fragment in which genomic regions of
alyA flanked a cassette conferring blasticidin resistance to
transformed cells. As soon as clones appeared on the primary
Petri dish, we re-cloned them by limited dilution in liquid
growth medium. In the first attempt to isolate a mutant lacking alyA, we screened about 100 clones by PCR analysis of
genomic DNA and obtained a single mutant cell line. Cell
homogenates from this clone 74 had a lysozyme activity that
was reduced to about 50% of wild-type levels. When plated on
a bacterial lawn, the mutant initially formed plaques where the
food bacteria were consumed that grew only to half the size
measured for the wild-type strain. This phenotype, however,
was not stable. Plating experiments repeated over the course of
2 weeks revealed that the diameter of plaques increased in the
alyA mutant until no difference was seen to wild-type cells. In
contrast, the enzyme activity remained low.
10438
Compensation of Lysozyme Deficiency
FIG. 2. Sequence features of the alyA gene. Nucleotide sequence
of the cDNA clone SSF469 and derived amino acid sequence. The
residues identified by peptide sequencing are shown in boldface. CS
denotes the cleavage site positions between the signal peptide (residues
1–19), the mature protein, and the prodomain (residues 139 –181).
Above the nucleotide sequence, H2 denotes the single HincII site used
for insertion of the resistance cassette and the Myc epitope, respectively
(see Figs. 3 and 5). The positions of the six introns from the genomic
clone are numbered I1-I6. The underlined sequences indicate the position of primers used for PCR in Fig. 3.
Because of this phenotypic instability, we repeated the transformation to obtain another mutant. In the second attempt, one
mutant was found among about 150 clones analyzed. The following analysis focuses on this clone, 138. PCR analysis using
a primer pair that binds to both alyA and alyB genes (see
below) revealed both genes in genomic DNA from wild-type
cells, whereas only a product corresponding to alyB was detected in the mutant (Fig. 3A). The mutant carried the blasticidine resistance cassette precisely within the alyA locus (Fig.
3B). As seen previously in strain 74, the knock-out mutant
alyA138 had a total lysozyme activity corresponding to about
40% of wild-type cells and maintained this level over more than
2 months (Fig. 3C). When the plaque diameters of the alyA138
strain were measured and compared with wild-type values,
they were found to increase from 60% to 90% within about 10
days, steadily rising to 150% after 40 days and finally stabilizing at a constant value corresponding to 180% of wild-type cells
(Fig. 3D). Despite these changes, repeated PCR analyses did
not reveal any alteration at the alyA locus (data not shown). All
of the subsequent experiments were performed using cells from
this stage.
FIG. 3. Unstable phenotype in an alyA knock-out mutant. A,
PCR products from genomic DNA of wild-type cells (AX2) and alyA138
using the primers flanking introns 4 and 5 (see Fig. 2). The upper band
corresponding to the alyA gene is missing in the mutant. The remaining
band was cut out and sequenced and, thus, identified as a product of a
second isoform, the alyB gene. The intron positions are conserved, but
their length varies. Intron 4 is 90 bp in alyA and 81 bp in alyB. Intron
5 is 125 bp in alyA and 79 bp in alyB. Together, the sizes of alyA and
alyB introns amount to a 55-bp difference detected in the PCR reaction.
M, molecular mass standards. B, PCR using a primer corresponding to
a sequence coding for the N terminus of the signal peptide and a primer
that is located within the blasticidin resistance cassette only amplifies
a product from alyA138 mutant DNA. C, enzymatic activity of lysozyme
was measured in wild-type cells (wt, filled symbols, set to 100%) or the
alyA138 mutant (open symbols) at intervals after transformation, and
the turnover rates were plotted over time (days). It did not change over
the 2-month period shown and remained constant thereafter. D, plaque
diameter of the alyA138 mutants increased with time (days). Single
cells were seeded together with lawns of bacteria onto agar plates, and
the diameter of the zone cleared by phagocytosis was measured on day
5 after inoculation. Wild-type values were set to 100%.
To test whether the increased plaque size was due to an
AlyA-independent increase in the capacity to degrade bacteria,
we fed the cells with GFP-expressing bacteria and measured
the disappearance of their fluorescence signal. Unexpectedly,
the alyA138 mutant performed as efficiently as wild-type cells
(Fig. 4A), although it was still reduced in lysozyme activity. To
find out whether the increased plaque size was caused by an
increased efficiency of phagocytosis, we measured the internalization of an indigestible particle, fluorescently labeled yeast.
Indeed, the rate of particle uptake by alyA138 mutant cells was
almost 2-fold higher than the rate of wild-type phagocytosis
(Fig. 4B), providing a reasonable explanation for the increased
Compensation of Lysozyme Deficiency
10439
plaque diameter. Because the rate of fluid-phase uptake was
unaltered (Fig. 4C), the phenotype observed in the alyA138
mutant is not caused by a general change in the endocytic
properties of these cells.
Rescue of Phenotypic Defects—In the case of unstable phenotypes, it is particularly important to check whether the reexpression of the product corresponding to the disrupted gene
restores the phenotype to the wild-type-like character. To this
end we inserted the entire coding region of alyA including the
introns into an expression vector and produced stably transformed cell lines in the alyA138 mutant background. RT-PCR
analysis of various clones generally revealed a more intense
band corresponding to the alyA gene (data not shown), which
reflects the high copy number integration of the plasmid into
the genome. In clone 2.4 phagocytosis was reduced to approximately wild-type levels (Fig. 4B), plaque size returned to normal (Fig. 4D), and the lysozyme activity in homogenates was
also partially rescued (Fig. 4E).
Because we failed to produce a polyclonal antibody in
chicken, which was suitable for the localization of the enzyme
in cells, we resorted to analyzing the distribution of genetically
tagged AlyA proteins bearing an Myc epitope in the middle of
the mature enzyme or a GFP extension at its C terminus (Fig.
5A). In the former construct the enzyme prodomain was maintained, whereas the GFP tag eliminated the cleavage signal,
replacing the C-terminal prodomain. These constructs were
individually transformed into the alyA mutant background.
The presence of the transgene was verified by PCR (Fig. 5B),
and the protein was detected in Western blots (Fig. 5C). For all
rescued clones we established that the endogenous copy of alyA
was still disrupted (data not shown). Although expression of
each of the tagged lysozymes increased the enzymatic activity
of cell extracts only mildly (Fig. 5D), the efficiency of phagocytosis approached wild-type levels (Fig. 5E), indicating that the
hybrid molecules were sorted to proper destination in the cells.
Localization of the alyA Product—Both tagged proteins localized to numerous small vesicles distributed throughout the
cytoplasm. Because the GFP moiety replaced the prodomain in
the hybrid construct, we compared the subcellular distributions of AlyA-GFP and Myc-tagged AlyA directly in cells transformed with a mixture of both plasmids. Fig. 6A documents
that the majority of GFP-containing vesicles are also loaded
with the Myc-tagged enzyme. The perinuclear ring and the
peripheral structures labeled with the anti-Myc-antibody alone
represent the endoplasmic reticulum (data not shown), suggesting that the Myc epitope in the middle of the AlyA protein
reduces the efficiency of protein folding. To investigate the
identity of the labeled vesicles, we checked the AlyA-GFPexpressing strain for the distribution of three classes of lysosomal enzymes known to reside in different types of lysosomes.
FIG. 4. Endocytic performance of the alyA mutant and rescue
of the phenotype. A, degradation of bacteria by alyA138 cells (open
circles) in vivo occurs with the same efficiency as in the wild-type strain
(filled circles). Cells were challenged with GFP-expressing bacteria, and
the decrease of intracellular fluorescence was measured over time. B,
the rate of phagocytosis is increased in the alyA138 mutant and can be
rescued to wild-type-like levels upon re-expression of alyA (open
squares). Cells were incubated together with fluorescent yeast particles,
and the intracellular fluorescence signal (relative units) originating
from phagocytosed particles was measured at 15-min intervals. r.u.,
relative units. C, fluid-phase uptake is unaltered in the alyA138 mutant. Cells were allowed to internalize nutrient medium containing
fluorescently labeled dextran, and intracellular fluorescence (relative
units) was recorded over time. D, re-expression of AlyA reverts the
plaque size to wild-type-like dimensions. Conditions were as for Fig. 3D.
Filled bars are from wild-type cells, open bars represent the alyA138
mutant, and hatched bars denote the rescued strain. Values (in mm) are
the mean of 6 –7 measurements in at least three independent experiments (n as indicated). Error bars are S.E. r.u., relative units. E,
re-expression of alyA increases total enzyme activity about 2-fold. Enzymatic activity in cell extracts is measured as the decrease in absorbance of a bacterial cell wall preparation over time.
10440
Compensation of Lysozyme Deficiency
FIG. 5. Tagged AlyA proteins can complement the phenotypic
alterations of the alyA138 mutant. A, schematic drawing of the tagged
constructs. signal indicates the N-terminally cleaved signal peptide for
translocation into the endoplasmic reticulum. The active mature enzyme
is followed by the C-terminally cleaved prodomain (pro). Myc and GFP
tags were inserted in the middle of the mature protein (catalytic) or
replacing pro, respectively. B, PCR of Myc and GFP rescues produced in
the alyA138 background. In clones bearing the Myc epitope within the
coding sequence (Myc) the PCR-fragment was up-shifted in size relative to
the alyA band from wild-type genomic DNA (AX2) because the construct
was based on the genomic clone. In cells expressing the GFP-hybrid
protein (GFP) the PCR fragment was smaller than the genomic alyA
product because in this case the fusion protein was constructed using the
cDNA clone. Marker (M) sizes are in kilobases. C, Western blot of Myctagged and GFP-tagged proteins. Two membranes were incubated with
anti-Myc and anti-GFP monoclonal antibodies, respectively, and developed with a chromogenic substrate. Molecular masses of the tagged proteins correspond to the expected sizes of 15 and 40 kDa, respectively. The
asterisk indicates a cross-reactive band. The molecular masses of the
markers (M) are in kDa. D, rescue of enzymatic activity by Myc and GFP
total lysozyme activity in cell extracts (measured as in Fig. 4E) is increased by expression of Myc-tagged protein (open triangle) or by a GFP
hybrid (open squares). Wild-type cells (filled circles) and the alyA138
mutant (open circles) were measured as controls. E, rescue of phagocytosis
by expression of Myc- and GFP-tagged proteins. Particle uptake was
measured as in Fig. 4B. Symbols are used as in D.
FIG. 6. Localization of Myc- and GFP-tagged AlyA. Shown are
individual channels and overlay images of single confocal sections
through AlyA-GFP expressing cells (green) stained with various antibodies in indirect immunofluorescence (red). A, a cell expressing additionally Myc-tagged AlyA as detected by mAb 9E10. B, a cell stained for
common-antigen 1 using mAb 221–342-5. C, immunofluorescence using
mAb AD7.5 directed against the N-acetylglucosamine-1 phosphate
modification or the crystal protein, which is the target of mAb 130-80-2
binding (D). For E, cells were allowed to phagocytose latex beads as
seen in the transmission image. Bar, 5 ␮m.
Surprisingly, neither enzymes bearing the common-antigen 1,
a mannose-6-SO4-containing oligosaccharide (Fig. 6B), nor enzymes characterized by the N-acetylglucosamine 1-phosphate
modification co-distribute with AlyA (Fig. 6C). Vesicles containing crystal-forming esterases are also devoid of AlyA-GFP
(Fig. 6D). We, therefore, conclude that the enzyme resides in a
novel type of vesicle in Dictyostelium. To test whether these
vesicles deliver their contents into phagosomes, AlyA-GFPexpressing cells were fed with synthetic particles. One of the
rare events, where internalized beads are surrounded by a rim
of fluorescence, is shown in Fig. 6E. More frequently we saw
fluorescent granules accumulating in the vicinity of a bead as if
docked to the phagosome membrane (also visible in Fig. 6E).
We assume that the protein is rapidly degraded when it encounters the degradative enzymes of phagosome but is rather
stable in the storage vesicles.
Compensation of Lysozyme Deficiency
DISCUSSION
The only lysozymes of protozoan origin characterized at the
molecular level to date are from E. histolytica (20, 46), a parasitic amoeba colonizing the human small intestine. Entamoeba is thought to have diverged from the evolutionary path to
higher organism virtually at the same time as Dictyostelium
(47), and it was tempting to think that Dictyostelium lysozymes
are more closely related to the Entamoeba lysozymes than to
enzymes from other organisms. We purified the major protein
with lysozyme activity from extracts of Dictyostelium amoebae.
The protein was characterized and N-terminally sequenced,
and a subsequent data base search identified a gene product
with unknown function and allowed the elucidation of the
entire primary structure of the protein. The enzyme described
here is not related to any known lysozyme, and we suggest
viewing it as the founding member of a new lysozyme class.
Several other sequences for putative gene products that may
have lysozyme-like properties can be extracted from the Dictyostelium data base (Fig. 7A). A recent survey of the genome
reveals the existence of 11 lysozyme genes that potentially code
for lysozymes belonging to four classes; two are of the bacteriophage T4 type, and three are homologous to the C-type lysozymes. Dictyostelium also has two genes that may code for
proteins closely related to the unconventional lysozymes first
described from E. histolytica (20, 46). Searching with the sequence of the AlyA protein turns up three additional members
of this so far unique lysozyme gene family. The predicted protein sequences of ALY B and ALY C are characterized by 12
and 22 exchanged residues, respectively. The fourth and most
divergent member of the proteins encoded by the aly family,
ALY D, carries a 77-residue insertion in the middle of the
molecule, which consists almost exclusively of serines and glycines. The remaining sequence is only about 40% identical to
AlyA. In summary, no other organism is as well equipped with
lysozyme genes as Dictyostelium.
Many of the animals investigated so far appear to possess a
single type of lysozyme only; multiple isoforms of a single
lysozyme type can be found when the protein has also been
recruited as a digestive enzyme (23). The only other organisms
known to date bearing genes potentially coding for more than
one type of lysozyme are D. melanogaster and C. elegans. They
possess c-type and i-type lysozymes or E. histolytica-type and
i-type lysozymes, respectively (16, 46). Phylogenetic analysis
revealed that the newly recognized AlyA-D described here represent a novel type of lysozymes (Fig. 7B). It is reasonable to
assume that the amoebic lysozymes, as in other phagocytic
cells, are constituents of an intracellular armament that fulfill
a digestive function to exploit bacteria for nutrition. In addition, these enzymes may prevent uncontrolled microbial
growth within the digestive vacuoles.
Irrespective of whether AlyA is tagged with a Myc epitope in
the middle of the molecule or with a GFP moiety replacing the
C-terminal prodomain, the enzyme accumulates in the same
type of small vesicle (Fig. 6A). This observation indicates that
the prodomain is not required for proper sorting of AlyA, an
observation that has been made with other lysosomal enzymes
as well (48, 49). Because the N-terminal signal peptide is likely
to be cleaved immediately after entry into the endoplasmic
reticulum, the mature enzyme must encompass all the signals
required for correct subcellular targeting.
Interestingly, AlyA is concentrated in a specific sort of vesicle
devoid of other lysosomal enzymes (Fig. 6). This finding increases the number of primary lysosomes to four, because it has
been convincingly shown that mannose-6-SO4-modified enzymes and N-acetylglucosamine 1-phospate-modified proteins
10441
do not coexist in the same vesicle (39), and the number and
morphology of esterosomes (40, 50) identifies the AlyA-containing organelles as a novel class of lysosomes. A possible reason
for sorting AlyA away from the majority of proteases lies in the
sensitivity of the molecule; it is very short-lived in total cell
extracts at acidic conditions and becomes increasingly stable
during purification. Indeed, immunofluorescence experiments
reveal that bead-bearing phagosomes only rarely contain detectable steady state levels of the enzyme (Fig. 6), but its acidic
pH optimum is in good agreement with the pH values found
within Dictyostelium endosomes.
When alyA, the gene encoding the major lysozyme isoform, is
disrupted, the total lysozyme activity of cells extracts is reduced to 40% of wild-type cells as measured by the degradation
of bacterial cell walls in vitro (Fig. 3C). In support of these
findings, the initial ability of cells to form plaques on a lawn of
bacteria is reduced to a similar degree (Fig. 3D). Taken together these two observations clearly support a role for AlyA in
efficient degradation of phagocytosed bacteria, which is perfectly consistent with its well known activity to cleave the
bacterial peptidoglycan, the major constituent of the cell wall of
Gram-positives. As peptidoglycan is also a constituent of the
cell wall of Gram-negative bacteria, it is not surprising to find
the same initial results when alyA mutant cells grow on lawns
of E. coli and K. aerogenes.
However, when the alyA-deficient cells are cultivated for prolonged times, a surprising observation can be made. The efficiency of plaque formation increases gradually until it reaches
wild-type levels, finally exceeding these values by almost 2-fold
(Fig. 3D). The reasons for these phenotypic changes are not
trivial, because the enzymatic activity of lysozyme in both strains
remains constantly at 40% of wild-type level in vitro (Fig. 3C).
Most importantly, alyA mutant cells attain wild-type properties
after re-expression of wild-type and hybrid AlyA transgenes
(Figs. 4 and 5), indicating that the changes observed in the
mutant over time are not the consequence of an accumulation of
a number of second site mutations. Instead, the compensatory
events leading to increased phagocytosis in the mutant are more
likely to involve metabolic changes, regulation within networks
of gene expression, or epigenetic phenomena.
In principle, compensation that leads to the increased plaque
size in the Dictyostelium alyA mutants can be of two kinds.
Either the ability to degrade bacteria is enhanced or the rate of
phagocytosis is augmented. The first possibility can be ruled
out because when the mutants were measured for their ability
to attack GFP-expressing bacteria, they perform similar to
wild-type cells (Fig. 4A). On the contrary, when the uptake of
non-degradable yeast particles is measured, the rate of phagocytosis for all strains corresponds to the behavior observed in
the plaque-forming assay (Figs. 4 and 5).
The rate of phagocytosis depends on the dynamics of the
actin cytoskeleton, and many Dictyostelium mutants affected
in actin-binding proteins show a reduced rate of particle uptake
(51). Interestingly, a number of mutants exist that are characterized by an increased rate of phagocytosis (52–54). When the
expression levels of a variety of these proteins were analyzed by
Western blotting of mutant cell extracts, no differences to the
wild type were detected (data not shown). On the other hand, it
may be informative to look in more detail into signal-transducing cascades. In particular, overexpression of the small
GTPases RacC or Rap1 is known to result in increased phagocytosis, whereas fluid-phase uptake remains unaffected (55,
56). Although these proteins could possibly signal to the cytoskeleton, it remains to be investigated how they would perceive the level of AlyA within the compartments of the secretory and/or endocytic pathway.
10442
Compensation of Lysozyme Deficiency
FIG. 7. Multiple sequence alignment and phylogenetic tree of lysozymes. A, multiple sequence alignment using ClustalX. The amino acid
sequences of the ALY isoforms A–D are shown as proenzymes. The other amino acid sequences of putative lysozymes of D. discoideum, namely
c-type (DdLysC1–3), phage-type (DdLysT41–2), and Entamoeba-type lysozymes (DdLysEh1–2), were found in databases and are aligned with
representative homologues of the respective class of lysozymes from other organisms. B, phylogenetic tree of lysozymes from different species
drawn and analyzed with PAUP 4.0. Data on sequences are given under “Experimental Procedures.” An unrooted distance tree (neighbor joining)
is presented. Numbers at the nodes represent bootstrap proportions on 1000 replicates; values greater than 70% are depicted only. The ALY family
members are marked by bold letters. The invertebrate lysozymes are considered a monophyletic family, and therefore, the entire branch is shaded.
Compensation of Lysozyme Deficiency
Acknowledgments—We thank Friedrich Buck for protein sequencing,
Thomas Jacobs for mass spectrometry, and Josef Rüschoff and
Thomas Brodegger for making their DNA sequencing facility available.
Günther Gerisch, Annette Müller-Taubenberger, and Hudson Freeze
have kindly provided monoclonal antibodies. We are grateful to all
teams involved in the Dictyostelium genome and cDNA sequencing
projects.
REFERENCES
1. Rybin, V., Ullrich, O., Rubino, M., Alexandrow, K., Simon, I., Seabra, M. C.,
Goody, R., and Zerial, M. (1996) Nature 383, 266 –269
2. Novick, P., and Zerial, M. (1997) Curr. Opin. Cell Biol. 9, 496 –504
3. Barbosa, M. D. F. S., Ngyuyen, Q. A., Tchernev, V. T., Ashley, J. A., Detter,
J. C., Blaydes, S. M., Brandt, S. J., Chotai, D., Hodgman, C., Solari, R. E.,
Lovett, M., and Kingsmore, S. F. (1996) Nature 382, 262–265
4. Winchester, B., Vellodi, A., and Young, E. (2000) Biochem. Soc. Trans. 28,
150 –154
5. Walkley, S. U. (2001) Curr. Opin. Neurol. 14, 805– 810
6. Buczynski, G., Bush, J., Zhang, L., Rodriguez-Paris, J., and Cardelli, J. (1997)
Mol. Biol. Cell 8, 1343–1360
7. Bruckert, F., Laurent, O., and Satre, M. (2000) Protoplasma 210, 108 –116
8. Harris, E., Wang, N., Wu, W., Weatherford, A., De Lozanne, A., and Cardelli,
J. (2002) Mol. Biol. Cell 13, 656 – 669
9. Cornillon, S., Dubois, A., Bruckert, F., Lefkir, Y., Marchetti, A., Benghezal, M.,
De Lozanne, A., Letourneur, F., and Cosson, P. (2002) J. Cell Sci. 115,
737–744
10. Saftig, P., Hetman, M., Schmahl, W., Weber, K., Heine, L., Mossmann, H.,
Koster, A., Hess, B., Evers, M., and von Figura, K. (1995) EMBO J. 14,
3599 –3608
11. Tyynela, J., Sohar, I., Sleat, D. E., Gin, R. M., Donnelly, R. J., Baumann, M.,
Haltia, M., and Lobel, P. (2000) EMBO J. 19, 2786 –2792
12. Journet, A., Chapel, A., Jehan, S., Adessi, C., Freeze, H., Klein, G., and Garin,
J. (1999) J. Cell Sci. 112, 3833–3843
13. Markart, P., Korfhagen, T. R., Weaver, T. E., and Akinbi, H. T. (2004) Am. J.
Respir. Crit. Care Med. 169, 454 – 458
14. Markart, P., Faust, N., Graf, T., Na, C. L., Weaver, T. E., and Akinbi, H. T.
(2004) Biochem. J. 380, 385–392
15. Jollès, P., and Jollès, J. (1984) Mol. Cell. Biochem. 63, 165–189
16. Bachali, S., Jager, M., Hassanin, A., Schoentgen, F., Jollès, P., Fiala-Medioni,
A., and Deutsch, J. S. (2002) J. Mol. Evol. 54, 652– 664
17. Watts, D. J., and Ashworth, J. M. (1970) Biochem. J. 119, 171–174
18. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H.,
Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk,
D. C. (1985) Anal. Biochem. 150, 76 – 85
19. Schägger, H., and von Jagow, G. (1987) Anal. Biochem. 166, 368 –379
20. Jacobs, T., and Leippe, M. (1995) Eur. J. Biochem. 231, 831– 838
21. Osserman, E. F., and Lawlor, D. P. (1966) J. Exp. Med. 124, 921–952
22. Shugar, D. (1952) Biochim. Biophys. Acta 8, 302–309
23. Dobson, D. E., Prager, E. M., and Wilson, A. C. (1984) J. Biol. Chem. 259,
11607–11616
24. Miller, G. L., and Golder, R. H. (1950) Arch. Biochem. Biophys. 29, 420 – 423
25. Reissig, J. L., Storminger, J. L., and Leloir, L. F. (1955) J. Biol. Chem. 217,
959 –966
26. Yamada, H., and Imoto, T. (1981) Carbohydr. Res. 92, 160 –162
27. Andrä, J., Berninghausen, O., Wülfken, J., and Leippe, M. (1996) FEBS Lett.
385, 96 –100
10443
28. Adachi, H., Hasebe, T., Yoshinaga, K., Ohta, T., and Sutoh, K. (1994) Biochem.
Biophys. Res. Commun. 205, 1808 –1814
29. Nellen, W., Datta, S., Reymond, C., Sivertsen, A., Mann, S., Crowley, T., and
Firtel, R. A. (1987) Methods Cell Biol. 18, 67–100
30. Faix, J., and Dittrich, W. (1996) FEBS Lett. 394, 251–257
31. Drengk, A., Fritsch, J., Schmauch, C., Rühling, H., and Maniak, M. (2003)
Curr. Biol. 13, 1814 –1819
32. Gerisch, G., Albrecht, R., Heizer, C., Hodgkinson, S., and Maniak, M. (1995)
Curr. Biol. 5, 1280 –1285
33. Hanakam, F., Albrecht, R., Eckerskorn, C., Matzner, M., and Gerisch, G.
(1996) EMBO J. 15, 2935–2943
34. Maniak, M., Rauchenberger, R., Albrecht, R., Murphy, J., and Gerisch, G.
(1995) Cell 83, 915–924
35. Hacker, U., Albrecht, R., and Maniak, M. (1997) J. Cell Sci. 110, 105–112
36. Maselli, A., Laevsky, G., and Knecht, D. A. (2002) Microbiology 148, 413– 420
37. Crameri, A., Whitehorn, E. A., Tate, E., and Stemmer, W. P. C. (1996) Nat.
Biotechnol. 14, 315–319
38. Neuhaus, E. M., Horstmann, H., Almers, W., Maniak, M., and Soldati, T.
(1998) J. Struct. Biol. 121, 326 –342
39. Souza, G. M., Metha, D. P., Lammertz, M., Rodriuez-Paris, J., Wu, R., Cardelli,
J. A., and Freeze, H. H. (1997) J. Cell Sci. 110, 2239 –2248
40. Bomblies, L., Biegelmann, E., Döring, V., Gerisch, G., Krafft-Czepa, H., Noegel, A. A., Schleicher, M., and Humbel, B. M. (1990) J. Cell Biol. 110,
669 – 678
41. Munro, S., and Pelham, H. R. (1987) Cell 48, 899 –907
42. Heringa, J. (1999) Comput. Chem. 23, 341–364
43. Swofford, D. (1999) PAUP (Phylogenetic Analysis Using Parsimony and Other
Methods), Sinauer Associates, Inc., Sunderland, MA
44. Morio, T., Urushihara, H., Saito, T., Ugawa, Y., Mizuno, H., Yoshida, M.,
Yoshino, R., Mitra, B. N., Pi, M., Sato, T., Takemoto, K., Yasukawa, H.,
Williams, J., Maeda, M., Takeuchi, I., Ochiai, H., and Tanaka, Y. (1998)
DNA Res. 5, 335–340
45. Glöckner, G., Eichinger, L., Szafranski, K., Pachebat, J. A., Bankier, A. T.,
Dear, P. H., Lehmann, R., Baumgart, C., Parra, G., Abril, J. F., Guigo, R.,
Kumpf, K., Tunggal, B., Cox, E., Quail, M. A., Platzer, M., Rosenthal, A.,
and Noegel, A. A. (2002) Nature 418, 79 – 85
46. Nickel, R., Jacobs, T., and Leippe, M. (1998) FEBS Lett. 437, 153–157
47. Bapteste, E., Brinkmann, H., Lee, J. A., Moore, D. V., Sensen, C. W., Gordon,
P., Durufle, L., Gaasterland, T., Lopez, P., Muller, M., and Philippe, H.
(2002) Proc. Natl. Acad. Sci. U. S. A. 99, 1414 –1419
48. Huete-Perez, J. A., Engel, J. C., Brinen, L. S., Mottram, J. C., and McKerrow,
J. H. (1999) J. Biol. Chem. 274, 16249 –16256
49. Moin, K., Demchik, L., Mai, J., Duessing, J., Peters, C., and Sloane, B. F.
(2000) Adv. Exp. Med. Biol. 477, 391– 401
50. Yuan, A., and Chia, C. P. (2000) Exp. Cell Res. 261, 336 –347
51. Maniak, M. (2002) Int. Rev. Cytol. 221, 257–287
52. Wienke, D. C., Knetsch, M. L. W., Neuhaus, E. M., Reedy, M. C., and Manstein,
D. J. (1999) Mol. Biol. Cell 10, 225–243
53. Temesvari, L., Zhang, L., Fodera, B., Janssen, K. P., Schleicher, M., and
Cardelli, J. A. (2000) Mol. Biol. Cell 11, 2019 –2031
54. Konzok, A., Weber, I., Simmeth, E., Hacker, U., Maniak, M., and MüllerTaubenberger, A. (1999) J. Cell Biol. 146, 453– 464
55. Seastone, D., Lee, E., Bush, J., Knecht, D., and Cardelli, J. (1998) Mol. Biol.
Cell 9, 2891–2904
56. Seastone, D., Zhang, L., Buczynski, G., Rebstein, P., Weeks, G., Spiegelman,
G., and Cardelli, J. (1999) Mol. Biol. Cell 10, 393– 406