Microtubule-assisted mechanism for functional metabolic macromolecular complex formation

Abstract

Evidence has been presented for a metabolic multienzyme complex, the purinosome, that participates in de novo purine biosynthesis to form clusters in the cytoplasm of living cells under purine-depleted conditions. Here we identified, using fluorescent live cell imaging, that a microtubule network appears to physically control the spatial distribution of purinosomes in the cytoplasm. Application of a cell-based assay measuring the rate of de novo purine biosynthesis confirmed that the metabolic activity of purinosomes was significantly suppressed in the absence of microtubules. Collectively, we propose a microtubule-assisted mechanism for functional purinosome formation in HeLa cells.

Enzymes synthesizing inosine monophosphate through a de novo purine biosynthetic pathway (Fig. 1A) have long been hypothesized to form a multienzyme complex in cells (1–3). Our investigation of this hypothesis in vivo successfully revealed that the human de novo purine biosynthetic enzymes colocalize in the cytoplasm of human cell lines upon purine depletion (Fig. 1 B and C) (4, 5). Subsequently, we proposed a subcellular metabolic organization for de novo purine biosynthesis, the “purinosome,” in cells (4). More importantly, the association and dissociation of the purinosome was regulated by changing the purine levels or by manipulating the activity or expression levels of protein kinase CK2 in live cells (4, 5).

Fig. 1.

Cellular localization of hFGAMS-GFP participating in de novo purine biosynthesis. (A) De novo purine biosynthetic pathway transforms phosphoribosyl pyrophosphate (PRPP) to inosine monophosphate (IMP) in 10 steps. AIRS, aminoimidazole ribonucleotide synthetase; AICAR Tfase, aminoimidazole carboxamide ribonucleotide transformylase; ASL, adenylosuccinate lyase; CAIRS, carboxyaminoimidazole ribonucleotide synthase; FGAMS, formylglycinamidine ribonucleotide synthase; GAR, glycinamide ribonucleotide synthetase; GARS, GAR synthetase; GAR Tfase, GAR transformylase; IMPCH, IMP cyclohydrolase; PPAT, PRPP amidotransferase; and SICARS, succinylaminoimidazole carboxamide ribonucleotide synthetase. Steps 2, 3, and 5 are catalyzed by a trifunctional enzyme, TrifGART; steps 6 and 7 are catalyzed by a bifunctional enzyme, PAICS; and steps 9 and 10 are catalyzed by a bifunctional enzyme, ATIC. (B and C) Distribution of hFGAMS-GFP transiently expressed in HeLa cells grown in purine-rich (B) and purine-depleted (C) media. (Scale bar, 10 μm.)

Because cytoskeletal structures have been proposed to play an important role in the organization of metabolic enzymes (3), we explored, in this work, whether the purinosome is associated with cellular structural elements. For example, glycolytic enzymes including aldolase were identified as bound to actin cytoskeleton in mammalian and yeast cells (6–8). Interestingly, dynamic alternation of actin structures during the cell cycle seems to be correlated with the glycolysis-mediated production of ATP to satisfy an increased demand for energy (9). Therefore, we sought the structural and functional relationships between the purinosome and cellular cytoskeletal structures using human formylglycinamidine ribonucleotide synthase (hFGAMS) fused with monomeric green/orange fluorescent proteins (GFP/OFP) as a purinosome marker.

To visualize or manipulate the cytoskeleton in the presence of purinosomes, we probed cellular actin networks using rhodamine-conjugated phalloidine to stain F-actin structures within fixed cells and alternatively inhibited actin polymerization by the addition of cytochalasin D into live cells. In parallel, we stained microtubule filaments in live cells using a TubulinTracker Green reagent (Taxol conjugated with Oregon Green 488) and also treated live cells with nocodazole, which directly binds to tubulin so as to interfere with microtubule formation in cells. Moreover, we established a cell-based assay monitoring the flux of de novo purine biosynthesis to demonstrate the metabolic functionality of purinosomes in the presence and the absence of small molecules. Collectively, we propose that the spatial distribution of functionally active purinosomes is controlled by the network of microtubules when cells demand purine production.

Results

Microtubule-Associated Purinosome Formation.

We investigated whether cytoskeletal structures are involved in the purinosome assembly. First, we stained actin filaments with rhodamine-phalloidine in fixed HeLa cells and also microtubules with a TubulinTracker Green reagent in live HeLa cells in the presence of hFGAMS-GFP/OFP as a purinosome marker. The cytosolic clusters did not colocalize with the actin network (Fig. 2). However, purinosomes were found associated with microtubule filaments in the cytoplasm (Fig. 3). In addition, we examined how purinosome assembly responds to the small-molecule inhibitors cytochalasin D and nocodazole, which interfere with actin and microtubule polymerization, respectively. The addition of cytochalasin D in living HeLa cells did not have an impact on the distribution of purinosomes (Fig. 4 A and B). However, it was apparent that the dissociation of purinosomes occurred upon the addition of nocodazole (Fig. 4 C and D). These data with small molecules are consistent with the cytoskeleton staining experiments described above. Collectively, depolymerization of microtubules by nocodazole appears to disfavor the cluster formation of purinosomes even under purine-deficient conditions.

Fig. 2.

Localization of purinosomes and actin filaments in fixed HeLa cells grown in purine-depleted medium. (A) Transiently expressed and subsequently fixed hFGAMS-GFP–forming clusters in the cytoplasm (green channel). (B) Actin networks stained by rhodamine-phalloidine in the cytoplasm (red channel). (C) Merged image of hFGAMS-GFP (A, green in C) and actin cytoskeleton (B, red in C). (Scale bar, 10 μm.)

Fig. 3.

Subcellular localization of purinosomes harbored by microtubule filaments in HeLa cells grown in purine-depleted medium. (A) Microtubule networks stained by a TubulinTracker Green reagent in the cytoplasm (green channel). (B) Transiently expressed hFGAMS-OFP–forming clusters in the cytoplasm, representing formation of purinosomes (red channel). (C) Merged image of microtubules (A, green in C) and hFGAMS-OFP (B, red in C). (D) Representative region of interest highlighted in the white box in panel (C). Of note, the enlarged image of panel (D) was enhanced for clarification by adjustments of brightness, contrast and/or color balance. (Scale bar, 10 μm.)

Fig. 4.

Effects of small molecules on purinosome assembly formed in HeLa cells grown in purine-depleted medium. Cytochalasin D (A and B) or nocodazole (C and D) was supplied to HeLa cells displaying purinosomes formed by hFGAMS-GFP. Individual images were taken before addition of the inhibitors (untreated; A and C) and after the cells had been incubated with the inhibitors for a given time (B, 90 min; D, 60 min). (Scale bar, 10 μm.)

Suppression of Purinosome Activity by Nocodazole.

We then established a cell-based assay to monitor the flux of de novo purine biosynthesis in the presence and the absence of inhibitors. Lawns of HeLa cells were pulsed with [¹⁴C(U)]-glycine, a substrate of glycinamide ribonucleotide synthetase at step 2 of de novo purine biosynthesis. Its incorporation into purines, the rate of which represents the flux of de novo purine biosynthesis, was determined via acid extraction and ion exchange resin column chromatography. The ¹⁴C incorporation into purines was normalized to the total number of cells in the assay, plotted versus the time after the pulse, and showed its linear incorporation as a function of time.

Without an inhibitor, the rate of ¹⁴C incorporation of glycine into the pool of cellular purines in HeLa cells grown under purine-depleted conditions was ∼42% greater than the rate observed for cells grown under purine-rich conditions (Fig. 5A). We then treated cells with the microtubule disrupting agent nocodazole to assess its effect on the functionality of purinosomes. After a 1-h incubation with nocodazole, [¹⁴C(U)]-glycine was similarly pulsed for 3 h (Fig. S1). Although purine-rich HeLa cells barely responded to nocodazole (Fig. 5B), the flux of de novo purine biosynthesis for purine-depleted HeLa cells was suppressed by ∼36% at the 3-h time point in the presence of nocodazole relative to the DMSO control (Fig. 5B). This experiment clearly showed that purine-deficient cells had diminished de novo purine biosynthesis in the absence of microtubules. Thus, HeLa cells maintained in purine-depleted conditions rely on the functional clustering of purinosomes that is spatially organized by microtubule networks.

Fig. 5.

Metabolic flux measurement of de novo purine biosynthesis for HeLa cells. (A) De novo purine biosynthesis is measured by determining amount of [¹⁴C(U)]-glycine incorporated into cellular purines in HeLa cells cultured in purine-rich (■) and purine-depleted (□) media. Incorporation was found to be linear with time up to 4 h, and ratio of de novo purine biosynthesis rates in purine-depleted to purine-rich media was 1.42 by fitting data with the least-squares line method (10, 11). However, data could alternatively be fit with a single exponential function, resulting in larger difference between the two data sets (i.e., 1.60). Error bar indicates SD of three independent assays. Of note, the data points at t = 0 from the two cell culture conditions overlap. (B) Effects of nocodazole on de novo purine biosynthesis was evaluated in a similar way by measuring [¹⁴C(U)]-glycine incorporation for 3 h (Fig. S1). For each type of cells, de novo purine biosynthesis was compared in the absence (i.e., DMSO) and presence of nocodazole. Purine biosynthesis in purine-depleted HeLa cells was decreased by ∼36% in the presence of nocodazole at 3 h. Bar height is the ¹⁴C incorporation into purines per million cells. Error bar indicates SD of three independent assays. *Unpaired one-tailed Student t test revealed that the effect of nocodazole on purine-depleted HeLa cells was statistically significant (P < 0.001). It should be noted that the cells for Fig. 5A were maintained in the preferred growth medium until harvesting, whereas the cells for Fig. 5B were rinsed with buffered saline solution to be treated with nocodazole and then maintained in buffered saline solution until harvesting, to be consistent with cellular imaging conditions.

Discussion

We conducted cellular imaging of cytoskeletal structures in the presence of purinosomes. Purinosomes were clearly embedded within a network of microtubules, but actin filaments were not associated or colocalized with purinosomes. In addition, disruption of the microtubule network by the addition of nocodazole was sufficient to dissociate purinosomes in live HeLa cells. The spatial distribution of purinosomes results from their being embedded within microtubule networks.

We then demonstrated that the association and dissociation of purinosomes were correlated with the rate of de novo purine biosynthesis in HeLa cells. By monitoring ¹⁴C-glycine incorporation into the pool of cellular purines, we were able to observe increased purine biosynthesis in cells with purinosomes. More importantly, nocodazole attenuated the metabolic flux of de novo purine biosynthesis in purine-depleted HeLa cells. Therefore, this cell-based purinosome activity assay indeed supports the role of microtubules for functional purinosome formation in live cells.

The observed localization phenotype between purinosomes and microtubules (Fig. 3) is also supported by our previous observations. When a region of interest containing a purinosome cluster was photobleached, fluorescent intensities were recovered in the same location as the photobleached area (4). In addition, in experiments in which sequential enrichment and depletion of purine levels triggered the dissociation and the association of purinosomes (4), reclustering of purinosomes did not occur in the same location. Therefore, we propose that newly forming purinosomes would stochastically nucleate at any location in the cytoplasm guided by microtubules but would remain at that location until they functionally dissociate.

As mentioned earlier, glycolytic enzymes are reversibly bound to actin structures, and glycolysis-driven ATP production was associated with actin cytoskeleton dynamics (9). In parallel, the anticipated association of de novo purine biosynthesis with de novo ATP synthesis suggests an advantage for the subcellular localization of purinosomes to microtubules owing to their need for a ready energy source to perform microtubule-mediated cellular dynamics. Alternatively, because purinosomes are distributed across the entire cytoplasm, microtubule-assisted functional purinosome formation might be an alternative means of maintaining cellular energy homeostasis throughout the cytoplasm. Although it is possible that specific microtubule-associated proteins facilitate purinosome formation, we conclude that the network of microtubules minimally provides nucleation sites for functionally active purinosome formation in the cytoplasm upon purine starvation.

Materials and Methods

Materials.

The hFGAMS-GFP and hFGAMS-OFP constructs were prepared as described before (4). Rhodamine-phalloidine and TubulinTracker Green were purchased from Molecular Probes. Cytochalasin D and nocodazole were obtained from Sigma. [¹⁴C(U)]-Glycine was from DuPont/New England Nuclear.

Transfection of Mammalian Cells.

A human cervical cancer cell line, HeLa (ATCC), was maintained and transfected for this study as described before (4). Briefly, HeLa cells were subjected to the following: “purine-depleted medium,” RPMI 1640 (Mediatech) supplemented with dialyzed 5% FBS (Atlanta Biological) and 50 μg/mL gentamicin sulfate (Sigma); and “purine-rich medium,” MEM (Mediatech) with 10% FBS and 50 μg/mL gentamicin sulfate. FBS was dialyzed against 0.9% NaCl at 4 °C for ∼2 d. Lipofectamine 2000 (Invitrogen) as a transfection reagent was used by following the manufacturer's protocol as previously described (4). Of note, an alternative purine-depleted medium (i.e., MEM, dialyzed 10% FBS and 50 μg/mL gentamicin sulfate) was evaluated with respect to purinosome formation in HeLa cells by transiently expressing hFGAMS-GFP.

Fluorescence Microscopy of Live and Fixed Cells.

All samples were imaged at ambient temperature (∼25 °C) with a 60× objective (1.49 numeric aperture; Nikon Apo TIRF) using a Photometrics CoolSnap ES² CCD detector mounted onto a Nikon TE-2000E inverted microscope as described before (5). Oregon Green 488 and GFP detection was accomplished using a S484/15x excitation filter (Chroma Technology), S517/30m emission filter (Chroma Technology), and Q505LP/HQ510LP dichroic (Chroma Technology). Rhodamine and OFP detection was carried out using a S555/25x excitation filter (Chroma Technology), S605/40m emission filter (Chroma Technology) and Q575LP/HQ585LP dichroic (Chroma Technology).

Cytochalasin D and nocodazole were added to cells after three washes with buffered saline solution (20 mM Hepes, pH 7.4, 135 mM NaCl, 5 mM KCl, 1 mM MgCl₂, 1.8 mM CaCl₂ and 5.6 mM glucose). Cells transiently expressing hFGAMS-GFP were imaged before and after the addition of either 2 μL cytochalasin D (1 mg/mL in DMSO) or 4 μL nocodazole (4.2 mg/mL in DMSO) to give final concentrations of 1 μg/mL cytochalasin D and 8 μg/mL nocodazole, respectively. Control experiments were also performed by the addition of 4 μL DMSO.

To stain cellular microtubule filaments, live HeLa cells transiently expressing hFGAMS-OFP were washed with buffered saline solution, followed by incubation with a TubulinTracker Green reagent (1 mM in DMSO; final concentration, 250 nM) at 37 °C for 30 min. In addition, to investigate the cellular distribution of actin filaments in fixed HeLa cells, cells were prepared similarly to those used for live cell imaging; however, the cells transfected with hFGAMS-GFP were fixed with freshly prepared 3% formaldehyde, permeabilized with 0.2% Triton X-100, and blocked with 10% normal goat serum (Jackson ImmunoResearch Laboratory) for 30 min at RT as described before (4). The cells were then incubated for 20 min at RT with rhodamine-phalloidine (8.3 μM in DMSO; 5 μL/sample) in PBS (PBS: 10 mM Na₂HPO₄, pH 7.4, 2 mM KH₂PO₄, 137 mM NaCl, and 2.7 mM KCl).

Determination of de Novo Purine Biosynthetic Rates.

The rate of de novo purine synthesis was determined by the incorporation of [¹⁴C(U)]-glycine (DuPont/New England Nuclear, NEC-276E, 111.70 mCi/mmol) into cellular purines using the method of Boss and Erbe (10). HeLa cells were maintained in purine-rich and purine-depleted media (MEM supplemented with 10% FBS or 10% dialyzed FBS, respectively, with 50 μg/mL gentamicin sulfate) for at least three passages. These cells were then seeded into T75 flasks containing the appropriate, gentamicin-free media at 2 × 10⁶ and 3 × 10⁶ cells/flask for the purine-rich and purine-depleted conditions, respectively. After allowing ≈36 h for the cells to achieve midlog phase growth, the cells were placed in 2 mL fresh media. After reequilibration, the cells were pulsed with [¹⁴C(U)]-glycine (125 μM, 20 mCi/mmol, 5 μCi/flask) for the desired time. The media was aspirated, and the cells were washed three times with 10 mL ice-cold Dulbeccos's PBS (Cellgro). Cells were harvested by treatment with 0.25% trypsin–EDTA solution.

To each cell pellet, 1 mL perchloric acid (0.4 M) was added, followed by vigorous vortexing to suspend cells. Incubation of cell suspensions in a boiling water bath for 1 h completely lysed the cells and extracted all purines. Immediately after the acid extraction, the tubes were cooled on ice. Cellular debris was pelleted by centrifugation and the supernatant was loaded onto 0.8 × 3 cm AG50W-X8 (100–200 mesh, Bio-Rad) columns that had been preequilibrated with 0.1 M HCl. The columns were washed with 5 mL HCl (1 M), and purines were then eluted with 5 mL of HCl (6 M). Quantitation was achieved by mixing 1 mL of the eluant with 10 mL Ecoscint (National Diagnostics) followed by liquid scintillation counting using a Beckman Coulter LS6500 instrument. To obtain comparable rates between purine-rich and purine-depleted conditions, de novo purine biosynthesis was normalized to the total number of cells.

Effects of Nocodazole on de Novo Purine Biosynthetic Rate.

HeLa cells cultured in purine-rich and purine-depleted media were inoculated into T75 flasks following the protocol for measuring the de novo purine biosynthetic rate. On the day of assay, cells were rinsed and equilibrated in buffered saline solution for 1 h before adding nocodazole (final concentration, 8 μg/mL) or DMSO as control. After an additional 1 h incubation with nocodazole or DMSO, the cells were pulsed with [¹⁴C(U)]-glycine (125 μM, 20 mCi/mmol, 5 μCi/flask) and incubated at 37 °C for 1, 2, and 3 h before harvesting the cells to measure ¹⁴C incorporation into purines as described above. ¹⁴C incorporation into newly synthesized purines was normalized to the total number of cells. We also performed an unpaired one-tailed Student t test using Microsoft Excel to determine whether the effect of nocodazole on purine-depleted HeLa cells was statistically significant.