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First published online 8 July 2008
doi: 10.1242/jcs.026682

Journal of Cell Science 121, 2473-2480 (2008)
Published by The Company of Biologists 2008
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Research Article

The secreted Dictyostelium protein CfaD is a chalone

Deenadayalan Bakthavatsalam, Debra A. Brock, N. Neda Nikravan, Kevin D. Houston, R. Diane Hatton and Richard H. Gomer*

Department of Biochemistry and Cell Biology, MS-140, Rice University, 6100 S. Main Street, Houston, TX 77005-1892, USA

* Author for correspondence (e-mail: richard{at}rice.edu)

Accepted 30 April 2008


   Summary
Top
 Summary
Introduction
 Results
 Discussion
 Materials and Methods
 References
 
Dictyostelium discoideum cells secrete CfaD, a protein that is similar to cathepsin proteases. Cells that lack cfaD proliferate faster and reach a higher stationary-phase density than wild-type cells, whereas cells that overexpress CfaD proliferate slowly and reach the stationary phase when at a low density. On a per-nucleus basis, CfaD affects proliferation but not growth. The drawback of not having CfaD is a reduced spore viability. Recombinant CfaD has no detectable protease activity but, when added to cells, inhibits the proliferation of wild-type and cfaD– cells. The secreted protein AprA also inhibits proliferation. AprA is necessary for the effect of CfaD on proliferation. Molecular-sieve chromatography indicates that in conditioned growth medium, the 60 kDa CfaD is part of a ~150 kDa complex, and both chromatography and pull-down assays suggest that CfaD interacts with AprA. These results suggest that two interacting proteins may function together as a chalone signal in a negative feedback loop that slows Dictyostelium cell proliferation.

Key words: Autocrine factor, Chalone, Cell number counting, Tissue size


   Introduction
Top
 Summary
 Introduction
Results
 Discussion
 Materials and Methods
 References
 
Secreted factors called chalones inhibit the proliferation of the secreting cells, forming a negative feedback loop that can regulate the number of the cells secreting the chalone (Gamer et al., 2003Go; Gomer, 2001Go). As the number of cells that secrete the chalone increases, the concentration of the chalone and the associated inhibition of proliferation increases. An example of a chalone is the polypeptide myostatin, which is made by, and secreted from, muscle. As the percentage of the body occupied by muscle increases, the serum concentration of myostatin increases (Lee and McPherron, 1999Go). Myostatin inhibits myoblast proliferation, and this negative feedback maintains the amount of muscle in the body (Thomas et al., 2000Go).

For many tissues, the identity of the associated chalone is unclear. In the phenomenon of tumor dormancy, tumors appear to secrete factors that inhibit the proliferation of metastatic cells (Cameron et al., 2000Go; Guba et al., 2001Go; Luzzi et al., 1998Go). Despite the potential use of such factors to inhibit the proliferation of metastases, these factors are largely unknown.

An excellent system to study secreted factors such as chalones is the simple eukaryote Dictyostelium discoideum (Kessin, 2001Go). Dictyostelium cells normally exist as haploid amoebae that eat bacteria on soil and decaying leaves; laboratory strains can also proliferate in a bacteria-free nutrient broth. When the amoebae starve, they cease to divide and begin to secrete an 80 kDa glycoprotein called conditioned medium factor (CMF). When there is a high density of starving cells, as indicated by a high concentration of CMF (Jain et al., 1992Go; Yuen et al., 1995Go), the cells aggregate using relayed pulses of extracellular cAMP as a chemoattractant (Aubry and Firtel, 1999Go). The aggregating cells form streams that break up into groups of ~20,000 cells (Shaffer, 1957Go). Each group develops into a fruiting body consisting of a mass of spore cells supported on a ~1-mm-high column of stalk cells.

A secreted ~450 kDa protein complex that is called counting factor (CF) modulates adhesion and motility during aggregation to regulate stream break-up, and thus group- and fruiting-body size (Brock and Gomer, 1999Go; Gao et al., 2004Go; Jang and Gomer, 2005Go; Roisin-Bouffay et al., 2000Go; Tang et al., 2002Go). We found that AprA, a 60 kDa protein in a partially purified CF preparation, is not a CF component but, rather, is part of a ~150 kDa complex that inhibits proliferation and, thus, has the properties of a chalone (Brock and Gomer, 2005Go). Here, we show that another protein, CfaD, is also not a component of CF. Instead, CfaD is part of a ~150 kDa complex, interacts with AprA and, similar to AprA, has the properties of a Dictyostelium chalone.


   Results
Top
 Summary
 Introduction
 Results
Discussion
 Materials and Methods
 References
 
CfaD is a cathepsin-L like protein but lacks the protease activity
Some preparations of partially purified CF contained a 27 kDa protein. The amino acid (aa) sequence of a tryptic peptide of this protein matched part of an open reading frame in the Dictyostelium genome (supplementary material Fig. S1). We named the predicted protein CfaD for CF-associated protein. The predicted molecular mass of CfaD is 58.6 kDa, suggesting that the 27 kDa protein is a breakdown fragment of CfaD. The predicted CfaD aa sequence contains a peptidase C1A motif and is, over a stretch of 315 aas, 34% similar to cathepsin L precursors from the mosquito Aedes aegypti and other species (supplementary material Fig. S2). Cathepsins are a family of proteases responsible for protein turnover in the lysosome (Nomura and Katunuma, 2005Go). Tumors often contain increased levels of cathepsins and, unlike normal cells, secrete cathepsins, which appear to promote invasion by degrading the surrounding extracellular matrix (Gocheva and Joyce, 2007Go; Jedeszko and Sloane, 2004Go).

CfaD also shows 34% similarity to the 26/29 kDa proteinase of the flesh fly Sarcophaga peregrine (supplementary material Fig. S2), which is synthesized as a ~62 kDa polypeptide with a 19-aa signal sequence. This protein can hydrolyse the cathepsin substrate Z-Phe-Arg-AMC (Fujimoto et al., 1999Go). During the processing of the 26/29 kDa proteinase, the signal sequence is removed and the remaining protein is cleaved into a 23 kDa and an ~25 kDa fragment, whereby the 13 kDa fragment of the precursor that lies between the 23 kDa and 25 kDa fragments is then discarded (Fujimoto et al., 1999Go). Both the 23 and 25 kDa subunits are post-translationally glycosylated, and the resulting 26 kDa and 29 kDa fragments are secreted by hemocytes into the hemolymph of larvae to degrade the larval midgut and fat body during metamorphosis (Fujimoto et al., 1999Go; Nakajima et al., 1997Go; Takahashi et al., 1993Go). In CfaD, there is a predicted 18-aa signal sequence, and the aa sequence of a tryptic peptide of the secreted form of CfaD begins at the predicted signal sequence cleavage site (supplementary material Fig. S1, arrow), suggesting that the secreted form of the 27 kDa fragment of CfaD (CfaD-27) begins with VPQL.

A comparison of the predicted CfaD aa sequence with other cathepsin sequences (Berti and Storer, 1995Go; Santamaria et al., 1998Go) indicated that CfaD contains two key active site residues, a glutamine at position 327 and a cysteine at position 333 (supplementary material Figs S1, S2). However, CfaD belongs to the peptidase C1 family, which includes proteins without peptidase activity (Rawlings and Barrett, 1993Go). Using the protease assay that showed that the Sarcophaga 26/29-kDa proteinase has a protease activity (Fujimoto et al., 1999Go), we observed that in PBM (roughly mimicking the extracellular environment), a human cathepsin-L control had activities of ~2.5 and ~3.1 nM Z-Phe-Arg-AMC hydrolyzed/hour/µg protein at 22°C and 37°C, respectively. We observed that rCfaD (recombinant CfaD containing a His tag), rHMCfaD (recombinant CfaD containing both His and Myc tags), and rHMCfaD-PM (rHMCfaD with Gln327 changed to Lys and Cys333 changed to Gly) had no detectable protease activity at either 22°C or 37°C, with a detection limit of 0.004 nM Z-Phe-Arg-AMC hydrolyzed/hour/µg protein. In addition, rCfaD had no detectable protease activity at pH 5.2, roughly corresponding to the pH within a lysosome (Aubry et al., 1993Go). As described below, all three of the rCfaD variants inhibit Dictyostelium cell proliferation, suggesting that CfaD acts as a signal despite its lack of detectable enzymatic activity. This is similar to what we had previously observed for two functional components of the Dictyostelium CF group size regulation signal, CF45-1 (which has similarity to lysozymes but no detectable lysozyme activity) and CF60 (which has similarity to acid phosphatases but little or no acid phosphatase activity) (Brock et al., 2003bGo; Brock et al., 2006Go).

CfaD regulates proliferation
To elucidate the function of CfaD, we disrupted cfaD by homologous recombination. A northern blot indicated that wild-type cells contain a 1.9 kb cfaD mRNA, and that there is no detectable cfaD mRNA in cfaD– cells (Fig. 1A). Affinity-purified anti-CfaD antibodies stained a 65 kDa and a 27 kDa band on western blots of total protein from mid-log phase wild-type cells, and these proteins were not detected in cfaD– cells (Fig. 1B), which suggests that both proteins are encoded by cfaD, and that the affinity-purified anti-CfaD antibodies are specific for CfaD. Compared with wild type, there were higher levels of both bands in the cells of the CfaD-overexpressing strain cfaDOE, similar levels in cfaD– cells that overexpressed CfaD (cfaD–/cfaDOE), and somewhat lower levels in cells of the aprA– strain (that do not express AprA) (Fig. 1B). In other experiments, the levels of CfaD and CfaD-27 (the 27 kDa band) were essentially the same in wild-type and aprA– cells (data not shown). We had observed previously that the levels of the CF component countin are variable when other CF components are missing (Brock et al., 2003bGo). Staining cells with affinity-purified anti-CfaD antibodies showed that all vegetative wild-type cells contain CfaD, whereas cfaD– cells do not show appreciable staining (supplementary material Fig. S3). Deconvolution microscopy indicated that CfaD is concentrated in subcellular structures, possibly vesicles (supplementary material Fig. S3).


Figure 1
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  		Fig. 1. cfaD– cells lack cfaD mRNA and CfaD protein. (A) Northern blot of RNA from wild type (WT) and cfaD– vegetative cells probed for cfaD. A loading-control gel stained with ethidium bromide showed apparently equal quantities and lack of degradation of the ribosomal RNA bands. (B) Western blot of total cell lysates from mid-log vegetative cells stained with affinity-purified anti-CfaD antibodies. Molecular weight markers (in kDa) are given at the right. A loading-control gel stained with Coomassie Blue showed apparently equal quantities of proteins in all samples. (C) Disruption of cfaD affects the appearance of fruiting bodies. Cells of the indicated strain were grown on bacterial lawns and fruiting bodies were photographed. Bar, 0.5 mm.

 

Compared with parental wild-type cells, the cfaD– cells formed large fruiting bodies with large spore heads, whereas cells overexpressing CfaD formed tall fruiting bodies (Fig. 1C). Expression of CfaD in the cfaD– cells caused these cells to form fruiting bodies that, compared with the cfaD– fruiting bodies, resembled wild-type fruiting bodies (Fig. 1C). Together, the data suggest that lack of, or overexpression of, CfaD affects development, and that to a first approximation, expression of CfaD in the cfaD– cells rescues the phenotype, suggesting that the phenotype of the cfaD– cells is due to disruption of cfaD.


Figure 2
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  		Fig. 2. CfaD slows the proliferation of cells growing in liquid shaking culture. (A) Cells were diluted to 2x105 cells/ml in HL5 and the cell density was measured daily. Values are the mean ± s.e.m. from six independent experiments. The absence of error bars indicates that the error was smaller than the plot symbol. WT, wild-type. At day 9, all of the cfaD– cells appeared to be dead. The saturation densities (in units of 107 cells/ml) were 2.4±0.1 for wild type, 3.6±0.3 for cfaD–, 1.5±0.1 for cfaDOE, and 2.2±0.1 for cfaD–/cfaDOE. The differences between all values are significant (P<0.05) except WT versus cfaD–/cfaDOE, which was not significant (1-way ANOVA, Tukey's test). (B) The data from the first 3 days were plotted using a log scale for the density. (C) Proliferation of cells growing on a lawn of bacteria was measured by plating 103 cells on bacteria and counting the number of cells at the indicated times. Values are the mean ± s.e.m. from three independent experiments.

 
Proliferation curves for cells growing in liquid shaking culture indicated that cfaD– cells proliferate faster than wild-type cells and reach stationary phase at a significantly higher cell density, whereas cfaDOE cells proliferate slower and reach stationary phase at a lower density (Fig. 2A,B). The cfaD–/cfaDOE cells, which have roughly as much CfaD as wild-type cells, showed proliferation rates and saturation densities that were roughly comparable with those of wild-type cells. From day 1 to day 3, the average doubling times were 12.4 hours for wild-type cells, 10.2 hours for cfaD– cells, 17.7 hours for cfaDOE cells, and 11.0 hours for cfaD–/cfaDOE cells (Fig. 2B). After reaching saturation density, cfaD– cells died off faster than wild type or cfaD–/cfaDOE cells, whereas cfaDOE cells were still alive when wild-type or cfaD–/cfaDOE cells had died (Fig. 2A). When grown on plates spread with bacteria, there were no significant differences in the proliferation of wild-type cells and the three transformants (Fig. 2C). By contrast, aprA– cells proliferate faster than wild-type cells on bacteria, suggesting that CfaD and AprA have different functions. The observed doubling times for wild-type cells in shaking culture and on bacteria were similar to those observed previously (Brock and Gomer, 2005Go; Loomis, 1982Go).

The absence of CfaD results in reduced spore viability
The evolutionary advantage for Dictyostelium to have AprA appears to be that, although it slows proliferation, it increases spore viability (Brock and Gomer, 2005Go). We observed that cells lacking CfaD also form structures with a reduced spore count and reduced spore viability, and that expressing CfaD in the cfaD– background partially rescues both defects (Table 1). In this and a previous report using this assay (Brock and Gomer, 2005Go), we observed that only ~1/3 of wild-type spores are viable, possibly due to the detergent used to wash the spores in the assay. Nonetheless, our results suggest that, like AprA, CfaD confers an evolutionary advantage to Dictyostelium cells because it increases spore viability.


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  		Table 1. CfaD affects spore viability

 

On a per nucleus basis, CfaD does not affect growth
Cells lacking AprA proliferate faster than wild-type cells, and tend to be multinucleate (Brock and Gomer, 2005Go). Compared with those of wild type, cfaD– cells also tended to be multinucleate, whereas cfaD–/cfaDOE cells had nuclei numbers similar to those in wild-type cells (Table 2). We did not observe wild-type or cfaD–/cfaDOE cells with more than four nuclei, whereas some cfaD– cells had as many as eight nuclei. This effect was also seen for cells growing in HL5 on a plastic surface (data not shown).


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  		Table 2. Effect of CfaD on the mass and protein content of cells

 

The growth (the increase in mass or protein per hour) and the proliferation (the increase in the number of cells per hour) of cells can be regulated independently (Dolznig et al., 2004Go; Gomer, 2001Go; Jorgensen and Tyers, 2004Go; Saucedo and Edgar, 2002Go). The absence of CfaD did not appear to affect mass or protein content of cells (Table 2). The values for wild-type cells are in agreement with previously reported values (Ashworth and Watts, 1970Go). After normalizing to the number of nuclei, on average cfaD– cells have less mass and protein per nucleus than wild-type or cfaD–/cfaDOE cells (Table 2). Since cells will roughly double their mass in one doubling time, a rough estimate of the growth rate can be obtained by dividing the cell mass or protein content by the doubling time. On a per-cell basis, cfaD– cells accumulate more mass and nuclei per hour than wild-type cells, but do not have a significantly higher protein accumulation (Table 3). When the growth was calculated per nucleus, there was no significant difference in the mass or protein accumulation rate between cfaD– and wild type (Table 3). Together, the data suggest that, although cells that lack CfaD have a shorter mitotic cycle, proliferate faster, and on a cell basis accumulate more mass per hour than do wild-type cells, the increased growth rate is due to the increased nuclear and cellular proliferation and is not due to an increased mass or protein accumulation per nucleus.


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  		Table 3. Effect of CfaD on the mass and protein increase of cells

 

CfaD interacts with AprA
CfaD-27 accumulates in conditioned growth medium from cells at ~5x105 cells/ml, a relatively low density, and then at densities above ~5x106 cells/ml CfaD-staining bands at 60, 55, and 37 kDa appear (Fig. 3B insert). All of these bands were absent in conditioned media from cfaD– cells, suggesting that the 60 kDa band is CfaD and the other bands are CfaD breakdown products (data not shown). Using known quantities of recombinant CfaD (Fig. 3A) as a standard, at a density of 1.2x107 cells/ml, there was ~84 ng/ml of CfaD, corresponding to ~7x10–6 ng CfaD per cell (Fig. 3B). Since the anti-CfaD antibodies are directed against the entire protein, we were unable to quantify the amount of CfaD-27.


Figure 3
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  		Fig. 3. The concentration of extracellular CfaD increases with cell density. (A) SDS-polyacrylamide gel of the rCfaD used for the calibration curve was stained with Coomassie Blue. Molecular mass markers (in kDa) are shown at left. (B) The amount of CfaD as a function of cell density was determined by staining western blots of conditioned growth medium from wild-type cells at the indicated densities together with standard amounts of rCfaD with affinity-purified anti-CfaD antibodies. Values are the mean ± s.e.m. from three independent experiments. The absence of error bars indicates that the error was smaller than the plot symbol. The insert shows a western blot of conditioned growth media harvested at the indicated densities (in units of 106 cells/ml) stained with affinity-purified anti-CfaD antibodies. Molecular mass markers (in kDa) are at left.

 
Molecular-sieve fractionation of conditioned growth medium and conditioned starvation medium followed by staining western blots of the fractions indicated that CfaD is present as a ~150 kDa complex and CfaD-27 as a ~115 kDa complex in both media (Fig. 4). We previously observed that wild-type-conditioned growth medium contains a broad ~150 kDa peak of activity that inhibits proliferation, and that this activity was not present in the conditioned growth medium from aprA– cells (Brock and Gomer, 2005Go). This suggests that the 150 kDa peak is the major peak of proliferation-inhibiting activity in wild-type cells.


Figure 4
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  		Fig. 4. Size fractionation of complexes containing CfaD and AprA. Conditioned starvation medium (CSM) and conditioned growth medium (CGM) from the indicated cells were concentrated, and were then fractionated using molecular-sieve chromatography. WT, wild type. Western blots of the different fractions were stained with affinity-purified anti-CfaD and anti-AprA antibodies. At the top, numbers indicate fraction number, and the position of molecular-sieve molecular-mass markers is indicated. A 670 kDa marker eluted at fraction 30. For each of the western blots, the position of molecular mass markers is indicated at left.

 
The loss of AprA decreases the apparent molecular mass of the CfaD-containing complex by 25 kDa and the CfaD-27-containing complex by 35 kDa (Fig. 4). However, these are less than the 60 kDa molecular mass of AprA. When using a column that had been used previously for several purifications, we had previously reported that in wild-type-conditioned starvation medium AprA eluted as a broad peak at ~150 kDa (Brock and Gomer, 2005Go). When using a new column of the same type, we observed a sharper peak at ~138 kDa (Fig. 4). The loss of CfaD decreases the apparent molecular mass of AprA by 30 kD, which is less than the molecular mass of CfaD, although similar to the molecular mass of CfaD-27. This suggests a physical link between AprA and CfaD. In addition, the apparent size of all of the complexes described above are smaller than that of the ~450 kDa CF, using the CF component CF60 as a marker (Brock et al., 2006Go) (and data not shown). This indicates that CfaD and CfaD-27 are not components of CF.

As an alternative way to determine whether there is an interaction between CfaD and AprA, we carried out pull-down assays. As shown in the upper left panel of Fig. 5, when rHMCfaD was added to either precleared wild-type- or aprA–-conditioned growth medium together with nickel beads, rHMCfaD was present in the pull-down samples (the material that bound to the nickel beads). When western blots of the pull-down samples were stained with anti-AprA (Fig. 5, bottom left panel), AprA was present in the samples from wild-type- but not aprA–-conditioned growth medium. This suggests that AprA binds to the rHMCfaD. Similarly, we were able to pull down CfaD by using rAprA (Fig, 5, right panels), further suggesting that AprA and CfaD interact with each other. Neither AprA nor CfaD were pulled down by the beads alone.


Figure 5
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  		Fig. 5. CfaD appears to interact with AprA. rHMCfaD and nickel-agarose beads were mixed overnight with conditioned growth medium (CGM) prepared from wild-type and aprA– cells. After washing with PBS, proteins bound to the beads were eluted with SDS sample buffer. Western blots of the bound proteins were stained with anti-Myc antibodies (top left panel) or anti-AprA antibodies (lower left panel). Similarly, rAprA was mixed with nickel-agarose beads and CGM from wild-type and cfaD– cells; western blots of the bound material were stained with anti-Myc antibodies (top right panel) or anti-CfaD antibodies (lower right panel). Molecular mass markers in kDa are shown in the middle.

 

The NC-4 strain of Dictyostelium secretes AprA and CfaD
The wild-type Dictyostelium strain used in these studies is an axenic strain derived from an isolate from North Carolina called NC4 (Sussman and Sussman, 1967Go). To determine whether NC4 cells also secrete AprA and CfaD, we grew NC4 cells on a lawn of bacteria on an agar plate, washed off the cells and bacteria, and analysed a solubilized part of the agar by western blotting (see Fig. 6A). As shown in Fig. 6A, NC4 cells secrete both AprA and CfaD into the agar. From 10 µl of agar, there was approximately 0.3 ng of CfaD (Fig. 6A). Measuring the diameter of and thickness of the agar in the plate, we can thus estimate that when there are ~3x107 cells on the plate, the agar contains ~850 ng CfaD. This then corresponds to an accumulation of 2.8x10–5 ng/cell, higher than the accumulation per cell for the axenic wild-type strain in shaking culture. When NC4 cells were grown in shaking culture with bacteria, the conditioned growth medium contained both AprA and CfaD, and molecular-sieve chromatography of this material showed a peak of both proteins at ~138 kDa (Fig. 6B). Together, the data suggest that CfaD and AprA are secreted by cells in the natural environment to slow proliferation.


Figure 6
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  		Fig. 6. NC4 wild-type cells secrete AprA and CfaD. (A) NC4 cells were grown on a bacterial lawn on an agar plate. The cells were washed off and a piece of the agar was mixed with SDS sample buffer, heated and, while still hot and molten, the material corresponding to 10 µl of the agar was loaded on gels. Western blots of the gels were stained with affinity-purified anti-AprA or anti-CfaD antibodies. A 0.5 ng recombinant CfaD standard (His-tagged rCfaD) was also loaded on the CfaD gel. Molecular mass markers in kDa are shown at left. (B) Size fractionation of medium conditioned by NC4 cells growing in shaking culture with bacteria, and western blotting of the fractions, was done as described for Fig. 4.

 

CfaD slows but does not stop proliferation
To test the hypothesis that CfaD acts as an extracellular signal that inhibits proliferation, recombinant CfaD was added to cells in growth medium. The recombinant CfaD inhibited the proliferation of wild-type and cfaD– cells (Fig. 7 and Table 4). CfaD appears to slow but not completely inhibit the proliferation of wild-type cells, because we observed that 640 ng/ml recombinant CfaD slowed proliferation to 75±2% of control. For unknown reasons, the maximal inhibition for cfaD– cells appears to be slightly less than that for wild-type cells. Recombinant CfaD had no observable effect on the proliferation of aprA– cells (Fig. 7), suggesting that AprA is necessary for the ability of extracellular CfaD to inhibit proliferation. CrlA has similarity with G-protein-coupled receptors, and crlA– cells proliferate faster than wild-type (Raisley et al., 2004Go). Recombinant CfaD inhibited the proliferation of crlA– cells, although the maximal inhibition was less than that for wild-type or crlA– cells, suggesting that CrlA is not necessary for the effect of CfaD on cells, but does potentiate its activity. For unknown reasons, the EC50 for recombinant CfaD to inhibit proliferation was lower in cfaD– and crlA– cells than in wild-type cells (Table 4). Fitting the data to a sigmoidal dose-response curve with a variable Hill coefficient gave a Hill coefficient of 1, indicating that there was no cooperativity in the dose-response curve. To determine whether mutating the putative cathepsin active site of CfaD affects the bioactivity of CfaD, we added rHMCfaD-PM to wild-type cells. At a final concentration of 150 ng/ml, rHMCfaD decreased proliferation at 12 hours by 23±4% (mean ± s.e.m., n=4), whereas rHMCfaD-PM decreased proliferation by 25±2%. Together, the data suggest that, CfaD acts as an extracellular signal that reduces cell proliferation, AprA is necessary for this effect, rHMCfaD and rCfaD have similar bioactivities, and the putative cathepsin-active site of CfaD is not necessary for its ability to slow proliferation.


Figure 7
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  		Fig. 7. Extracellular CfaD slows cell proliferation. His-tagged rCfaD at the indicated concentrations was added to cells growing in shaking culture, and cell densities were measured after 12 hours. For each experiment with each cell line, the proliferation at 12 hours was calculated as the density of cells treated with rCfaD as a percent of the density without rCfaD. Values are the mean ± s.e.m. from four separate experiments. The graphs show sigmoidal dose-response curves fit to the data; the calculated maximal inhibition and EC50 values for each cell line are given in Table 4.

 

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  		Table 4. Effect of recombinant CfaD on cells

 


   Discussion
Top
 Summary
 Introduction
 Results
 Discussion
Materials and Methods
 References
 
In this report, we found that CfaD is an autocrine secreted factor that slows cell proliferation, and thus has the properties of a chalone. At high levels of exogenous recombinant CfaD, the proliferation of wild-type cells was slowed by 31% over 12 hours. Given a 12.4 hour doubling time, this corresponds to changing the doubling time to ~25 hours. Similarly, the 23% decrease in the proliferation of cfaD– cells over 12 hours corresponds to changing the doubling time from 10.2 hours to ~16 hours. The increased doubling times roughly correspond to the observed 17.7-hour doubling time for cfaDOE cells.

There appear to be multiple secreted factors that slow Dictyostelium proliferation
Yarger et al. had described a secreted factor that inhibits proliferation at stationary phase (Yarger et al., 1974Go), has a molecular weight of less than 10 kDa and is heat stable. Since CfaD and AprA are large proteins, the AprA-CfaD complex is probably not identical with this factor. In addition, the factor described by Yarger appears at stationary phase and seems to completely stop proliferation, whereas we observed that AprA (Brock and Gomer, 2005Go) and CfaD only slow proliferation. This suggests that Dictyostelium cells use the AprA-CfaD complex to slow proliferation as the cells approach saturation, and use the factor described by Yarger to completely stop proliferation when cells reach stationary density.

CfaD and AprA also appear to have different properties. First, AprA inhibits the proliferation of cells growing on bacteria (Brock and Gomer, 2005Go), whereas CfaD does not. One explanation might be that AprA and CfaD interact with different receptors and signal transduction pathways, even though they are in the same complex. We previously observed this happening for countin and CF50, two components of the CF complex (Brock et al., 2003aGo; Brock et al., 2002Go). It is unclear why Dictyostelium cells would have a multi-protein chalone. The observation that recombinant CfaD does not slow proliferation when AprA is absent suggests that slowing proliferation requires the presence of both the AprA and the CfaD signal. Since Dictyostelium cells live in dirt, where there are presumably a large number of different compounds that could activate a receptor, one possibility is that having two different proteins function as signals would be the equivalent of having a message authenticator, decreasing the possibility of an exogenous compound `accidentally' triggering a decrease in proliferation at an inappropriate time.

Can slower proliferation be an advantage?
Compared with cells that contain CfaD, cells that lack CfaD proliferate faster but die faster in shaking culture and have reduced spore viability. One possible reason for the reduced viability of cfaD– cells is that the decreased amount of mass and protein per nucleus compared with wild-type cells represents less nutrients per nucleus; since protein synthesis per nucleus is not affected by the lack of CfaD, the cfaD– cells will thus run out of nutrients – especially amino acids – sooner than wild-type cells. The reduced spore viability might also be due to a similar reduction in the amount of nutrients per nucleus. Assuming that the key function of CfaD is to slow proliferation of vegetative cells, the advantage for Dictyostelium cells to use a chalone to slow proliferation would be increased fitness of cells when they are at densities where they may begin to starve.


   Materials and Methods
Top
 Summary
 Introduction
 Results
 Discussion
 Materials and Methods
References
 
Cell culture and molecular-sieve chromatography
Cell culture was done following Brock et al. (Brock et al., 1999) in HL5 medium (Formedium Ltd, Norwich, England) using wild-type Ax2 cells, aprA– strain DB60T3-8 (Brock and Gomer, 2005Go), and crlA– strain JH557 (Raisley et al., 2004Go). Conditioned growth and starvation media (CM) were prepared and concentrated, and PBM buffer was made, following Brock et al. (Brock et al., 2002Go). Size fractionation was carried out as described in (Brock and Gomer, 2005Go). Western blots of fractions were stained with anti-AprA antibodies as described in (Brock and Gomer, 2005Go). To examine the size of the CfaD complex secreted by NC4 cells, 1x106 NC4 cells were grown with live Klebsiella bacteria in PBM in a shaking suspension culture. As a control, bacteria were grown without Dictyostelium cells. After 36 hours, the Dictyostelium cell density was ~3x107 cells/ml, and the supernatant was clarified and used for gel filtration. Photography of aggregates and fruiting bodies was performed as described in Brock et al. (Brock et al., 2002Go). Proliferation assays, calculation of doubling times, staining of nuclei, spore counts and spore viability assays were done as described in (Brock and Gomer, 2005Go).

Disruption of cfaD
To generate a homologous recombination cfaD-knockout construct, PCR was performed using Ax2 genomic DNA as a template. All DNA fragments were ligated into pCR 2.1 (Invitrogen, San Diego, CA) and sequenced at Lonestar Labs (Houston, TX). PCR with the primers 5'-CGATAATCATCCGCCGGTATTAGGCCAAGCTCAC-3' and 5'-GCATGCTCTAGACCTGGGGTAGTGGTACAAACC-3' yielded a 1138 bp fragment of the 5' side of cfaD. This was digested with SacII and XbaI, and ligated into the same sites in pBluescript SK+ (Stratagene, La Jolla, CA) which had been previously modified to contain the 1.4 kb SmaI Cre-loxP blasticidin resistance cassette from pLPBLP (Faix et al., 2004Go) to generate pcf27/29-L. PCR was then carried out with 5'-GCAAATGTAAGCTTGTCTCGCCACCGAGTCCAAC-3' and 5'-CGCATTGGGCCCGGTTGGATATCAATCAAATCATTATC-3' to generate a 1056 bp fragment of the 3' end of cfaD. The fragment was digested with HindIII and ApaI and ligated into the same sites in pcf27/29-L to generate pcf27/29-LR. This was digested with SacII and ApaI, and the insert was purified by gel electrophoresis and a Geneclean II kit (Qbiogene, Inc., Carlsbad, CA). Dictyostelium Ax2 cells were transformed with the construct as described by Shaulsky et al. (Shaulsky et al., 1996Go). PCR and northern blot analysis were used to verify the disruption of cfaD. Seven cfaD disruption clones with the same phenotype were identified, and all of the results show data from clone DB27C-1, which is referred to in this report as cfaD–. RNA isolation and Northern blots were done following Brock et al. (Brock et al., 2002Go). The cDNA encoding the full-length secreted CfaD protein was used as a probe.

Expression of CfaD in Dictyostelium cells
To obtain a CfaD-overexpressing construct, PCR was carried out using a vegetative cDNA library and the primers 5'-GATACCGAGCTCATGAATAAATTCATTTTATTATTATC-3' and 5'-CAGCATCTCGAGTATTCTTTGTTGGAATTGG-3' to generate a fragment of the cfaD-coding region corresponding to the entire polypeptide starting with the first methionine, and a SacI site on one end and an XhoI site on the other to allow expression of a C-terminal Myc tag. After digestion with SacI and XhoI, the PCR product was ligated into the corresponding sites of pDXA-3D (Ehrenman et al., 2004Go) to produce the overexpression construct. Ax2 cells were transformed following Manstein et al. (Manstein et al., 1995Go), and expression of CfaD was verified by staining western blots of whole-cell lysates using anti-CfaD antibodies. The resulting CfaD-overexpressing strain was designated cfaDOE. Constructs were also made to express the 27 kDa and 29 kDa subunits separately, but only the full-length construct was successfully expressed in our hands. CfaD was similarly expressed in cfaD– cells, and the resulting strain was designated cfaD–/cfaDOE.

Preparation of recombinant His-tagged CfaD and antibody purification
Recombinant CfaD was prepared following the method used to prepare recombinant CF50 (Brock et al., 2002Go) with the exception that 5'-CTTATTCATATGGTTCCACAACTCCCAGCTGC-3' was used as the forward primer and 5'-CGGATCCTCGAGTTAATTCTTTGTTGGAATTGG-3' was used as the reverse primer for the PCR reaction. This resulted in a cDNA fragment encoding the region from the first aa of the putative secreted CfaD protein to the TAA stop codon. The resulting recombinant protein was designated rCfaD. Bethyl Laboratories (Montgomery, TX) used this protein to produce affinity-purified rabbit polyclonal anti-CfaD antibodies. Staining of western blots was carried out according to Brock et al. (Brock et al., 2002Go) using the affinity-purified anti-CfaD antibodies at 0.4 µg/ml.

Generation of His- and Myc-tagged rCfaD and rAprA
rCfaD with C-terminal His and Myc tags was generated in to facilitate pull-down assays. Following (Brock et al., 2002Go), the primers 5'-CTCGAGGTTCCACAACTCCCAGC-3' and 5'-TCTAGAGCATTCTTTGTTGGAATTGGATAGG-3' were used to generate a cfaD fragment corresponding to the secreted form of CfaD. This fragment was cloned into a TA cloning vector, pC2.1 (Invitrogen, Carlsbad, CA) which was then digested using XhoI and XbaI to cut out the cfaD fragment. The fragment was ligated into the XhoI and XbaI sites in pBAD/gIII(A) (Invitrogen) to construct pBAD-CfaD, with which Top-10 E.coli cells (Invitrogen) were transformed. To express the resultant protein (designated rHMCfaD), cells containing the pBAD-CfaD construct were grown overnight at 37°C in LB medium (Invitrogen). The overnight culture was then diluted with LB medium to an OD600 of 0.1 and further grown at 37°C. Once the culture reached an OD600 of 0.5, it was induced by adding 20% arabinose to a final concentration of 0.1%. After 5 hours of induction, cells were collected by centrifugation at 12,000 g for 15 minutes and resuspended in PBS (1.8 mM KH2PO4, 10.1 mM Na2HPO4, 137 mM NaCl, 2.7 mM KCl, pH 7.4) with EDTA-free protease inhibitors (Roche, Indianapolis, IN). The collected cells were disrupted using a cell disruptor (EmulsiFlex-05, Avestin, Canada) and 30% N-lauroylsarcosine sodium salt solution (Sigma Aldrich, St Louis, MO) was added to a final concentration of 5%. rHMCfaD was then purified using nickel-agarose beads (Qiagen, Valencia, CA) following the manufacturer's protocol. Using a similar protocol, rAprA was prepared using the primers 5'-CTCGAGATGGATTATGTCAATGCTCCTGAC-3' and 5'-GAATTCCAGTTGCAGTTGAACTAGCACT-3' to generate the expression plasmid pBAD-AprA.

Generating mutated rCfaD
Using pBAD-CfaD as a template, the primers 5'-CCCCAGTCAAAGATAAAGGTATTTGCGGTTCAGGTTGGACTTTTGG-3' and 5'-CCAAAAGTCCAACCTGAACCGCAAATACCTTTATCTTTGACTGGGG-3' were used in a PCR reaction to generate a mutated plasmid (pBAD-CfaD-PM) wherein Gln327 and Cys333 were replaced with Lys and Gly, respectively. The PCR reaction and transformation was carried out using a QuickChange Site-Directed Mutagenesis Kit (Stratagene, La Jolla, CA). The resulting plasmid pBAD-CfaD-PM was sequenced to confirm the two point mutations. The plasmid was then transformed into the Top-10 E.coli cells (Invitrogen) to express and purify the mutated recombinant rCfaD (designated rHMCfaD-PM) as described above.

Protease activity assay
An Innozyme cathepsin L activity kit (Calbiochem, La Jolla, CA) was used to determine the enzymatic activity of rCfaD, using either PBM or 50 mM MES (pH 5.2) as assay buffers. Human cathepsin L provided in the kit was used as a positive control and BSA as a negative control.

Pull-down assay
Wild-type and aprA– cells were grown, starting at 1x106 cells/ml, to a density of 5x106 to 6x106 cells/ml. After centrifugation at 3000 g for 3 minutes to clarify conditioned growth medium (CGM), 10 ml of CGM was passed through a 0.2 µm filter (Millipore, Bedford, MA) and was then concentrated to 1 ml using Amicon 10,000 MWCO ultra-filters (Millipore Corporation, Billerica, MA). To remove the proteins that non-specifically bind to nickel-agarose beads (Qiagen, Valencia, CA), the concentrated CGM was pre-cleared by mixing with 25 µl of nickel-agarose beads (50% slurry in PBS) at room temperature. After 1 hour, the pre-cleared CGM was collected by centrifugation at 17,500 g for 10 minutes. To 500 µl of pre-cleared wild-type or aprA– CGM, 1 µl of 500 ng/µl rHMCfaD was added together with 25 µl of nickel-agarose beads (50% slurry in PBS) and mixed end-to-end overnight at 4 °C. The beads were then washed five times with 1 ml of PBS and collected by centrifugation at 3000 g for 1 minute at room temperature. The beads were then collected using Zymo-P1 fast-spin columns (Zymos Research, Ontario, Canada) and material bound to the beads was eluted using 50 µl of 1xSDS sample buffer. The samples were heated at 95°C for 5 minutes and loaded onto 4-15% Tris-HCl gels (Biorad laboratories, Hercules, CA). Western blots of the gels were then stained either with a 1:1000 dilution of rabbit anti-Myc (Bethyl Laboratories, Montogomery, TX) or anti-AprA antibodies. Similarly, rAprA was used to pull down CfaD.

Proliferation inhibition assay
To assess the inhibitory effect of purified rCfaD on growing cells, cells were grown to 1x106 to 2x106 cells/ml, collected by centrifugation, and resuspended to 5x105 cells in 968 µl of HL5. Different concentrations of rCfaD in 32 µl of 20 mM potassium phosphate pH 6.4 were then added to the cells. After shaking for 12 hours, cells were counted. For each cell line, the percent proliferation was calculated as 100 x (cell count at 12 hours for a given rCfaD concentration) per (cell count at 12 hours with no CfaD added). A sigmoidal dose-response curve with a variable Hill coefficient of

Formula
was then fit to the data using nonlinear regression with Prism (GraphPad software, San Diego, CA).

Statistical analysis
Statistical analysis using GraphPad Prism software (GraphPad Prism software, San Diego, CA). Differences between two groups were assessed using the Mann-Whitney U test, or between multiple groups by ANOVA. Significance was defined as P<0.05.


   Acknowledgments
 
We thank Lisa Kreppel and Alan Kimmel for the gift of pLPBLP, Jeff Hadwiger and Dale Hereld for the crlA– cells, Darrell Pilling for helpful suggestions. This research was supported in part by the National Science Foundation under Grant No. PHY05-51164 to the Kavli Institute for Theoretical Physics, grant number C-1555 from the Robert A. Welch Foundation, and NIH GM074990.


   Footnotes
 
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/121/15/2473/DC1


   References
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 Summary
 Introduction
 Results
 Discussion
 Materials and Methods
 References
 

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