Fatty Acid Oxidation Is Required for Myxococcus xanthus Development

ABSTRACT

Myxococcus xanthus cells produce lipid bodies containing triacylglycerides during fruiting body development. Fatty acid β-oxidation is the most energy-efficient pathway for lipid body catabolism. In this study, we used mutants in fadJ (MXAN_5371 and MXAN_6987) and fadI (MXAN_5372) homologs to examine whether β-oxidation serves an essential developmental function. These mutants contained more lipid bodies than the wild-type strain DK1622 and 2-fold more flavin adenine dinucleotide (FAD), consistent with the reduced consumption of fatty acids by β-oxidation. The β-oxidation pathway mutants exhibited differences in fruiting body morphogenesis and produced spores with thinner coats and a greater susceptibility to thermal stress and UV radiation. The MXAN_5372/5371 operon is upregulated in sporulating cells, and its expression could not be detected in csgA, fruA, or mrpC mutants. Lipid bodies were found to persist in mature spores of DK1622 and wild strain DK851, suggesting that the roles of lipid bodies and β-oxidation may extend to spore germination.

IMPORTANCE Lipid bodies act as a reserve of triacylglycerides for use when other sources of carbon and energy become scarce. β-Oxidation is essential for the efficient metabolism of fatty acids associated with triacylglycerides. Indeed, the disruption of genes in this pathway has been associated with severe disorders in animals and plants. Myxococcus xanthus, a model organism for the study of development, is ideal for investigating the complex effects of altered lipid metabolism on cell physiology. Here, we show that β-oxidation is used to consume fatty acids associated with lipid bodies and that the disruption of the β-oxidation pathway is detrimental to multicellular morphogenesis and spore formation.

INTRODUCTION

Myxobacteria have provided fascinating insight into the evolution of social and emergent behaviors for well over a century. They are highly adaptable multicellular microorganisms that are ecologically widespread. Few species have been cultivated relative to the tremendous phylogenetic diversity observed by 16S ribosomal DNA sequencing. From the cultured species, several conclusions are obvious. First, their cell structure, macromolecular composition, and core metabolic processes resemble those of other members of the Deltaproteobacteria despite their unique morphological characteristics (1, 2). Second, their enormous genomes, often in excess of 10 Mbp, consist of single circular chromosomes filled with seemingly redundant genes (3, 4). Finally, their multicellularity is due to shared molecules that encourage common ventures. These molecules, both surface associated and extracellular, synchronize motility to facilitate cooperative or predatory behaviors depending on the circumstances (5–9).

The most visual standard of the myxobacterial life cycle is the migration of tens of thousands of cells to a common location, where they construct a fruiting body whose shape, size, and color vary among species. Inside the fruiting body, the cells differentiate into dormant environmentally resistant myxospores. Sporulation is an encystment process in which the entire cell shortens and rounds up to form a spore. In the case of Myxococcus xanthus, more than 80% of the membrane surface area is lost during spore formation (10).

Fruiting body development is induced by nutrient limitation and invigorated by the resourceful deployment of endogenous reserves. A striking cytological feature of developing M. xanthus cells is the appearance of lipid bodies composed of triacylglycerides (TAGs) (11). The sudden appearance of such an energy-rich resource in direct response to amino acid depletion represents a conundrum that we have investigated. Lipid bodies are formed from preexisting membrane phospholipids that are no longer necessary, as cells shorten during sporulation (10). The key step in this process appears to be the oxidation of cardiolipin and phosphatidylglycerol by the short-chain alcohol dehydrogenase CsgA (5). The unstable product undergoes multiple hydrations that fragment it into diacylglycerols (DAGs), orthophosphates, and dihydroxyacetone. From there, we suspect that one or more of the many annotated acyltransferases add the remaining fatty acids to convert these DAGs into TAGs.

The formation of lipid bodies offers appealing solutions both for cell shortening during spore formation and as a ready carbon and energy source for development. TAGs can be degraded into fatty acids and glycerol with lipases, which are abundant in M. xanthus (12). The most widespread and energetically favorable pathway for fatty acid utilization is β-oxidation, a cyclic pathway in which the fatty acid is shortened two carbons at a time to generate one molecule each of acetyl coenzyme A (acetyl-CoA), NADH, and reduced flavin adenine dinucleotide (FADH₂) with each turn of the cycle. The consumption of a single fatty acid can generate over 100 ATPs. The β-oxidation pathway is well characterized in Escherichia coli (13–18) and Pseudomonas (19–21). The core enzymes form an α2β2 complex encoded by fadB and fadA, respectively (14, 19, 22–25). During anaerobic β-oxidation, FadI and FadJ serve functions parallel to those of FadA and FadB in E. coli (26).

On the basis of microarray data, one set of M. xanthus genes homologous to those involved in fatty acid β-oxidation is upregulated during M. xanthus development (10, 27). These genes more closely resemble those found in anaerobic β-oxidation despite the fact that M. xanthus is an obligate aerobe. MXAN_5371, representing a fadJ homolog (new locus identifier [ID], MXAN_RS26065), is upregulated 4-fold, while MXAN_5372, representing a fadI homolog (new locus ID, MXAN_RS26070), is upregulated 2.4-fold after starvation for 18 h (10, 27). This time frame represents the peak of lipid body accumulation and is coincident with the onset of sporulation. Homologs MXAN_6987 and MXAN_6988, respectively (new locus IDs, MXAN_RS33820 and MXAN_RS33825, respectively), are not upregulated during development (10, 28).

In this work, we explored the hypothesis that β-oxidation is used to harvest the energy provided by lipid bodies during spore maturation. While the large number of redundant genes within the M. xanthus genome may have prevented the complete elimination of fatty acid utilization, our results suggest that β-oxidation is necessary for fruiting body development, spore coat synthesis, and spore resistance to heat and UV light.

RESULTS

Myxococcus xanthus contains two sets of β-oxidation genes.

Two genes predicted to be responsible for fatty acid degradation were upregulated during M. xanthus development at the peak of lipid body accumulation (10, 27). MXAN_5371, representing a fadJ homolog (new locus ID, MXAN_RS26065), was upregulated 4-fold. MXAN_5372, representing a fadI homolog (new locus ID, MXAN_RS26070), was upregulated 2.4-fold. These genes appear to be arranged in a two-gene operon, fadIJ. FadJ contains two domains (see Table S2 in the supplemental material). The first belongs to the crotonase/enoyl coenzyme A (enoyl-CoA) hydratase superfamily and the second resembles a short-chain alcohol dehydrogenase. Like the E. coli homologs, these domains could encode activities that perform successive steps in β-oxidation. A blastp search was conducted using the FadJ amino acid sequence (WP_011555336) with the M. xanthus protein database. Two other genes (MXAN_6987 [new locus ID, MXAN_RS33820] and MXAN_5136 [new locus ID, MXAN_RS24960]) appear to encode proteins that contain both hydratase and dehydrogenase domains. One of these genes, MXAN_6987, was highly similar to MXAN_5371, suggesting that a second copy of fadJ is present in M. xanthus. In addition, there were six other proteins homologous to the crotonase/enoyl-CoA hydratase superfamily domain of FadJ and two other proteins homologous to the dehydrogenase domain of FadJ (Table S2), which could also contribute to β-oxidation.

FadI (WP_011555337) is a putative thiolase, which performs the last step in β-oxidation. Results from the blastp search using this protein as a probe showed that there are four additional homologs, one of which (MXAN_6998 [new locus ID, MXAN_RS33825]) is in an operon with MXAN_6987, representing fadJ (Table S2). Only MXAN_5372 was upregulated during development (10, 27). Thus, it would appear that M. xanthus contains one set of fadIJ genes for development and a second for vegetative growth. To examine the roles of these genes, MXAN_5371 and MXAN_5372 were selected for markerless deletion, while MXAN_6987 was inactivated by plasmid insertion.

Deletion of fadI and fadJ results in a defective developmental phenotype.

The aggregation phenotype of M. xanthus cells during starvation-induced development was altered in the β-oxidation mutant strains LS4120 (MXAN_5371 and MXAN_5372 deletion) and LS4122 (LS4120 with a plasmid insertion in MXAN_6987) (Fig. 1, Table 1). During fruiting body morphogenesis, the wild-type strain DK1622 produces more aggregation foci than survive to become mature fruiting bodies. It selectively culls the population of smaller aggregates as aggregation proceeds (29, 30). In contrast, LS4120 and LS4122 produced greater abundances of fruiting bodies but were unable to eliminate many of the smaller aggregates as visualized in time-lapse movies of development. LS4120 and LS4122 also produced more irregularly shaped fruiting bodies than DK1622. This alteration in fruiting body morphogenesis did not result in a statistically significant change in the number of spores or viable spores produced by the mutant strains (Fig. 1). However, the spores produced by LS4122 were more susceptible to damage by heat and UV exposure (Fig. 2). Approximately 99% of LS4122 spores were no longer viable after exposure to 65°C for 60 min, while DK1622 spores lost viability to the same extent after 120 min (Fig. 2A). This same effect was not observed with LS4122 spores exposed to 60°C heat over the same time period (Fig. 2B). Exposure to UV irradiation at 1,000 μW/cm² for 90 s rendered >99.9% of LS4122 spores nonviable. By comparison, 150 s of UV irradiation exposure at this level was required to reduce DK1622 spore viability to <0.1% (Fig. 2C).

FIG 1.

Aggregation phenotypes of DK1622 and the β-oxidation pathway mutant strains LS4120 and LS4122 48 h poststarvation (top) and a comparison of sporulation and germination efficiencies of these strains (bottom). Direct spore counts are given as percentages of the wild-type strain DK1622. Viable spore counts are given as the percentage of each strain's direct spore counts. Spore counts were obtained from cells spotted onto TPM agar and incubated at 32°C for 5 days. Errors are the standard deviations (SDs) from two technical and three biological replicates.

TABLE 1.

Strains and plasmids used in this study

Strain or plasmid	Description	Reference or source
Strains
E. coli DH10B	Used for cloning of plasmids and protein expression	56
M. xanthus
DK1622	WT	3
DK851	Wild strain	57
LS3950	MXAN_5372 with Km insertion	This study
LS4120	MXAN_5371-MXAN_5372 in-frame deletion	This study
LS4122	MXAN_5371-MXAN_5372 in-frame deletion and MXAN_6987 with Km insertion	This study
LS4126	MXAN_6987 with Km insertion	This study
LS2442	ΔcsgA in-frame deletion	40
SW2808	mrpC in-frame deletion in DK1622	41
LS3600	fruA in-frame deletion	This study
LS4134	pHFS03 integrated in DK1622	This study
LS4136	pHFS03 integrated in LS2442	This study
LS4138	pHFS03 integrated in LS3600	This study
LS4140	pHFS03 integrated in SW2808	This study
Plasmids
pMR3487	M. xanthus 1.38-kb-PIPTG-MCS_A-PR4::lacI Tcr	58
pBJ113	M. xanthus markerless deletion plasmid	59
pCR2.1-TOPO	TA clone vector
pHFS01	pBJ113 containing MXAN_5371 and MXAN_5372 upstream and downstream flanking sequence	This study
pCRC4	pBJ113 containing MXAN_3117 upstream and downstream flanking sequences	This study
pHFS02	1,116-bp fragment of MXAN_6987	This study
pHFS03	M. xanthus MXAN_5372 UTR::tdTomato Tcr	This study

FIG 2.

Spore survival after exposure to 65°C heat stress (A), 60°C heat stress (B), or UV irradiation (C). Spore survival is expressed as a percentage of the spores plated for each time point. Percent survival values of <0.1 are given as 0.1. Error bars are SDs from two technical and two biological replicates.

During fruiting body formation, it has been observed that only about 20% of the cell population develop into spores, with the remaining 80% undergoing programmed cell death (31, 32). While the results presented here show a more dramatic decrease in rod-shaped cells within the first 24 h poststarvation, approximately 99%, the deletion of fadI and fadJ did not appear to alter the rate at which spores were produced or the extent of cell death during development (Fig. 3). These results indicate that cell fate determination is not noticeably affected by an interruption of the β-oxidation pathway. However, there may be a difference in the maturation of the mutant and DK1622 spores when examined at a finer level.

FIG 3.

Changes in the numbers of rod-shaped cells and spores over the course of development for M. xanthus DK1622 and the β-oxidation pathway mutant strains LS4120 and LS4122. Cells were spotted on TPM agar, harvested at the indicated time intervals, and counted in a Petroff-Hauser counting chamber. Error bars are SDs from two technical and four biological replicates.

fadJ and fadI removal increases lipid body content.

Following Nile red staining, the lipid body content was quantified in 30 cells for the triple mutant LS4122 and wild-type DK1622 cells, and the results are presented as a box plot (Fig. 4A). At 12 h, a fluorescence intensity of approximately 300 arbitrary units (AU) represents background levels of fluorescence due to Nile red binding to the phospholipids in the cell envelope. A standard t test revealed that the mutant had a statistically greater lipid content at 24, 36, and 96 h (Fig. 4A). This is particularly notable at 96 h, when many LS4122 rod-shaped cells still contained lipid bodies (Fig. 4B). However, the lipid body content still declined in the mutant.

FIG 4.

Lipid body production and retention in M. xanthus DK1622 and LS4122 cells during development. (A) Box plot of lipid body content based on fluorescence intensity of cells stained with the lipophilic Nile red dye. Each point indicates the fluorescence intensity of one cell. Horizontal lines inside the boxes indicate distribution medians. Tops and bottoms of each box indicate 75th (q₃) and 25th (q₁) percentiles, respectively. Whiskers extend to the highest and lowest points, or q₃ + 1.5(q₃ − q₁) and q₁ − 1.5(q₃ − q₁), whichever is closer to the median. P values were determined with a standard t test comparison of wild-type and mutant cells at each time point: 12 h, 0.08; 24 h, 0.00; 36 h, 0.00; 48 h, 0.89; 72 h, 0.56; 96 h, 0.00. (B) M. xanthus DK1622 and LS4122 cells stained with Nile red over the course of development. Scale bar, 5 μm. (C) M. xanthus DK1622 and LS4122 spores stained with Nile red 120 h into development. Scale bar, 5 μm.

The images of spores stained with Nile red at 120 h appear to contain lipid bodies, contradictory to a previous study which reported that lipid bodies have disappeared from DK1622 spores by 96 h (11). To verify this observation, spore thin sections were prepared at 48 h and 144 h for DK1622, LS4122, and DK851 and were examined by transmission electron microscopy (Fig. 5). These images show that lipid bodies are still prominent in all three strains at 144 h. The presence of lipid bodies in the wild strain DK851 indicates that the retention of lipid bodies is not unique to our lab strains. This result may reflect a more general carbon storage strategy for use during germination.

FIG 5.

Electron micrographs of thin sections of DK1622, LS4122, and DK851 spores during differentiation into spores. DK1622 spores at 48 h (A and B) and 144 h (E and F). LS4122 spores at 48 h (C and D) and 144 h (G and H). DK851 spores 144 h (I and J). Images of single spores were taken at ×12,000 magnification; scale bars, 0.2 μm. Images of groups of spores were taken at ×3,000 magnification; scale bars, 1 μm.

Deletion of fadI and fadJ slows spore coat maturation.

Electron micrographs of LS4122 and DK1622 spores taken at 48 h and 144 h show that the disruption of the β-oxidation pathway had a noticeable effect on the physical appearance of the spores (Fig. 5). LS4122 spores seemed to mature more slowly than DK1622 spores. The 144-h LS4122 spores appeared similar to the DK1622 48-h spores in terms of spore coat thickness and internal structure (Fig. 5A, B, G, and H). The spore coats present on DK1622 spores were on average twice the thickness of the spore coats on the LS4122 spores at both 48 h and 144 h poststarvation (Table 2). DK851 spores had spore coats of similar thickness to those of the LS4122 spores at 144 h (Table 2). Since DK851 was not included in any of the spore stress tolerance assays, it is not known if a thinner spore coat results in reduced spore viability in this strain. All spores were sonication resistant, regardless of variations in spore coat thickness. In addition to the large lipid bodies present in DK1622 and LS4122 spores, there were a number of smaller bodies that have not yet been characterized. These were not present in the 144-h-old DK1622 spores but were present in the LS4122 spores from this time point as well as in both strains' 48-h-old spores (Fig. 5). On the basis of these observations, LS4122 spores developed more slowly and less completely than DK1622 spores.

TABLE 2.

Spore coat thicknesses at 48 h and 144 h poststarvation for M. xanthus wild-type strain DK1622, β-oxidation pathway mutant LS4122, and the wild strain DK851

Strain	Thickness (μm) ata:
	48 h	144 h
DK1622	0.14 ± 0.04	0.28 ± 0.05
LS4122	0.06 ± 0.03	0.16 ± 0.03
DK851	NDb	0.14 ± 0.04

^aSpore coat thickness was calculated from electron micrographs. Values are ±SD from at least 20 spores.

^bND, no data collected.

MXAN_5372 expression occurs primarily in sporulating cells.

The 5′ untranslated region (UTR) of MXAN_5372 (fadI) was fused to tdTomato, resulting in strain LS4134. This transcriptional fusion was also created in the csgA (LS4136), fruA (LS4138), and mrpC (LS4140) deletion backgrounds. None of these deletion strains formed fruiting bodies or sporulated. The expression of the tdTomato fusion was analyzed at 0, 18, and 24 h by the quantification of fluorescence intensity in rod-shaped cells. MXAN_5372 expression was very low in vegetative cells (0 h) (Fig. 6). The expression reached the highest level in LS4134 after 18 h of starvation, in agreement with previous microarray results (27). In LS4136, LS4138, and LS4140, the tdTomato fusion maintained low levels of expression (Fig. 6). The fluorescence intensities of individual cells revealed a striking pattern when plotted against the cell lengths. The fluorescence intensity increased primarily in sporulating cells, which are roughly 1.5 to 4 μm in length (Fig. 7A). Fluorescence images of LS4134 cells also show that the fluorescence intensity of the spores is much more dramatic than that of rod-shaped cells (Fig. 7B). Together, these results argue that MXAN_5372 can be used as a spore-specific gene expression marker.

FIG 6.

Expression on the basis of fluorescence intensity of MXAN_5372 UTR::tdTomato in the DK1662 WT (LS4134), ΔcsgA (LS4136), ΔfruA (LS4138), or ΔmrpC (LS4140) backgrounds at 0 h, 18 h, and 24 h (54). Error bars are SDs from 30 cells selected to calculate the mean fluorescence intensity.

FIG 7.

Expression of MXAN_5372 UTR::tdTomato during development. (A) Relationship between MXAN_5372 expression level and cell length in LS4134 cells after 24 h of starvation. (B) Fluorescence images of LS4134 at different developmental time points.

FadJ and FadI mutants contain more FAD than DK1622.

We sought a method to directly quantify β-oxidation enzymatically. Initially, we tried to analyze the reverse FadJ dehydrogenase activity in cell extracts with acetoacetyl-CoA as the substrate and NADH as a cofactor (33). There was no difference between DK1622 and LS4122 (data not shown). As reported in other organisms and mitochondria, this is not surprising, as β-oxidation is not the only metabolic pathway to utilize acetoacetyl-CoA. Larger substrates more specific to β-oxidation are not commercially available, rendering this approach difficult.

Flavin adenine dinucleotide (FAD) levels are often used as a proxy for β-oxidation in mitochondria (34). FADH₂ is produced during the first step in β-oxidation and during the tricarboxylic acid (TCA) cycle. The TCA cycle genes encoding succinate dehydrogenase, the enzyme that produces FADH₂, are severely downregulated during development (27). Specifically, MXAN_3539 (MXAN_RS17155) and MXAN_3540 (MXAN_RS17160), which encode two succinate dehydrogenase subunits, are downregulated 8-fold (27). Since portions of the TCA cycle appear to be downregulated during development, this suggests that FADH₂ production at this stage would occur primarily from β-oxidation. Assuming that β-oxidation is the main source of FADH₂ during development, our mutants would then be expected to have higher levels of FAD. We quantified the FAD levels in DK1622 and in each of our mutants after 24 h of development. Strains LS4120 and LS4122 contained roughly 2-fold more FAD than DK1622 WT cells, in agreement with this principle (Table 3). These results suggest reduced rates of β-oxidation but are also consistent with the idea that the mutant strains are not completely deficient in β-oxidation.

TABLE 3.

FAD concentrations in M. xanthus DK1622 and β-oxidation pathway mutant strains

Strain	FAD concn (pmol/μg protein)a	MT/WTb
DK1622	0.36–0.45
LS4120	0.69–1.05	2.14
LS4122	0.67–1.04	2.12

^aReported as range of values from a single experiment with two replicates.

^bMT/WT, mutant FAD concentration divided by that of the wild type. Values are the averages from two replicates for each sample.

DISCUSSION

In this work, we examined the production and metabolism of lipid bodies by M. xanthus cells during fruiting body development. β-Oxidation is the most efficient pathway for fatty acid utilization. This pathway consists of a 4-step cycle involving oxidation of the fatty acyl-CoA, hydration, a second oxidation, and finally, thiolytic cleavage. The last three steps in fatty acid degradation are performed by a tetrameric complex consisting of two copies each of FadA and FadB in E. coli (15). The genes for these proteins are closely linked and may form an operon in E. coli (35). M. xanthus contains two sets of genes homologous to fadA and fadB, MXAN_5371 (annotated as FadJ) and MXAN_5372 (annotated as FadI), as well as MXAN_6987 and MXAN_6988. These M. xanthus homologs may also be arranged in operons.

M. xanthus development is induced by carbon limitation, and so the discovery that lipid bodies provide a large portion of the cell volume midway through development was surprising (11). Roughly 80% of the membrane surface area is lost during spore formation and used to produce lipid bodies containing TAGs. Bhat et al. proposed that M. xanthus development is composed of checkpoints that couple cell shortening to TAG/lipid body production to make the most effective use of the phospholipids that are no longer needed (10). The nature of the “factories” that couple the two processes is only beginning to be unraveled. A key enzyme involved in producing lipid bodies is CsgA (5). Long thought to be a protein morphogen (36–39), it is now clear that CsgA is an enzyme that creates DAGs from the phospholipids cardiolipin and phosphatidylglycerol. From there, the addition of the final acyl group creates the TAGs (5). Lipid body consumption ensures sufficient carbon and energy from TAG utilization for the production of spores. Concomitant with the appearance of the lipid bodies is the induction of a spore-specific β-oxidation pathway (MXAN_5371 and MXAN_5372) to consume the fatty acids. The highest MXAN_5372 promoter expression was seen in sporulating cells, indicating a requirement for β-oxidation in spore formation and/or germination. Increased expression of MXAN_5372 was not observed in mutant strains that do not form lipid bodies or sporulate (10, 40, 41). This includes mutants lacking the transcription factors MrpC and FruA as well as CsgA, all of which are essential for fruiting body formation and sporulation. Although the regulation of the MXAN_5372 promoter has not yet been examined, one possibility is that MrpC and FruA are essential for its expression.

The M. xanthus developmental program includes cells that follow different fates, including programmed cell death (∼80% of the initial population), peripheral rods that remain outside the fruiting bodies without sporulating (∼10%), and sporulating cells (∼10%) (42–45). The loss of β-oxidation genes did not change the overall rate of cell fate differentiation or the ratio of the different cell types. However, it did alter fruiting body morphogenesis, in that the mutants made more fruiting bodies and more irregularly shaped fruiting bodies. The mutants also did not remove smaller aggregates during development as is characteristic of DK1622. This result might suggest that β-oxidation is necessary for robust multicellular morphogenesis. Lipid bodies are first observed approximately 18 h after starvation and are commonly seen in both sporulating cells and cells undergoing programmed cell death but not in peripheral rods (6, 10, 11). However, lipid bodies seem to be unusually prevalent in peripheral rods from LS4122, where β-oxidation has been compromised by the loss of both canonical degradation pathways (Fig. 4, 96 h). It may be that β-oxidation is a property of all three cell types but to different extents. The β-oxidation pathway involving MXAN_6987 and MXAN_6988 is not upregulated during development.

The production of the rigid, compact spore coat is essential for the complete maturation of M. xanthus spores (46). A disruption of the spore-specific β-oxidation pathway in LS4122 results in 2-fold thinner spore coats than in DK1622. This structural deficiency is correlated with an increased susceptibility to heat and UV stress. The path of carbon flow can be deduced from this work in light of the upregulated genes in a published DNA microarray (27). Acetyl-CoA produced by β-oxidation enters the glyoxylate shunt, and from there, gluconeogenesis produces the monosaccharide substrates for spore coat formation. The proposed path of carbon flow is also consistent with results from previously published enzyme assays (47). The production of the spore coat requires a number of additional proteins, including a motor, transporters, and other proteins for the assembly of the coat components. The spore coat carbohydrates are transported to the cell surface by the outer membrane polysaccharide export machinery (Exo proteins) as the cell shortens (46, 47). These polysaccharides are then arranged by the Nfs complex of proteins into the tightly associated spore coat. The Agl motor is responsible for rotating the Nfs complex around the spore surface as it assembles the spore coat (46, 48, 49). The depletion of exogenous carbon and energy sources during starvation implies that M. xanthus must rely on internal carbon and energy stores, or stores released from lysed cells, to complete this process. Our results argue for the importance of β-oxidation given the thinner spore coats on β-oxidation pathway mutant spores and the susceptibility of the mutant spores to heat and UV stress.

Our results show that a portion of the lipid bodies remains within the mature spore. While it has been observed that some fatty acids are synthesized de novo during the germination of M. xanthus (50), we hypothesize that an impairment of β-oxidation would still result in fewer resources for metabolic activities needed during germination. Lipid bodies are the major energy resource for germination in plant seeds. Arabidopsis ped1 mutants, which lack a functional 3-ketoacyl-CoA thiolase, are consequently defective in fatty acid β-oxidation and require external sucrose to germinate (51). The residual lipid bodies maintained in mature spores could be used to provide energy during the early stages of germination. They could also be used to produce membrane phospholipids during outgrowth. Further studies will need to address the role of lipid bodies during germination.

MATERIALS AND METHODS

Bacterial strains and growth conditions.

The bacterial strains and plasmids used in this study are shown in Table 1. M. xanthus wild-type strain DK1622 and subsequent mutants were grown in charcoal-yeast extract (CYE) broth [1% Bacto Casitone, 0.5% yeast extract, 10 mM 3-(N-morpholino)propanesulfonic acid (MOPS), pH 7.6, and 4 mM MgSO₄] at 32°C with vigorous shaking. Cells were maintained on CYE supplemented with 1.5% Bacto agar. Kanamycin (Km) and tetracycline (Tc) were added to final concentrations of 50 μg · ml⁻¹ and 15 μg · ml⁻¹, respectively, when necessary. Development was induced by spotting cells on TPM agar plates (10 mM Tris HCl, 1 mM KH₂PO₄, 10 mM MgSO₄, 1.5% Difco agar, pH 7.6). E. coli was grown in Luria-Bertani (LB) medium (52).

Mutant construction.

The in-frame deletion of MXAN_5371 and MXAN_5372 was created in the DK1622 background by a double recombination method (53). Approximately 1,000 bp of DNA sequences flanking MXAN_5371 and MXAN_5372 were amplified by PCR. Primers MXAN_5371 plus 5372-UPF and MXAN_5371 plus 5372-UPR were used to amplify the upstream DNA fragment, and MXAN_5371 plus 5372-DOWNF and MXAN_5371 plus 5372-DOWNR were used for downstream DNA fragment amplification (see Table S1 in the supplemental material). The two DNA fragments were fused by overlapping PCR. The fused PCR product was digested with KpnI and HindIII and then cloned into the same sites of pBJ113 to create pHFS01 (Table 1). This plasmid was transformed into electrocompetent DK1622 cells. The initial recombination event was selected for on CYE with Km. After verifying that the resultant transformants were Km resistant, the secondary recombination event was selected for on CYE containing 1% galactose. The resulting strain, LS4120, was verified by PCR. The fruA (MXAN_3117 [new locus ID, MXAN_RS15110]) in-frame deletion strain, LS3600, was created using the same double recombination method. Primers FruA Upstream Fwd and FruA Upstream Rev were used to amplify a 1,000-bp region upstream of MXAN_3117, and primers FruA Downstream Fwd and FruA Downstream Rev were used to amplify a 1,000-bp region downstream of the target gene (Table S1). These PCR products were fused and used to create the deletion plasmid pCRC4, which was then transformed into DK1622 (Table 1). The selection for the recombination events was carried out as described above.

Strain LS4126 was generated via plasmid insertion in MXAN_6987. Primers MXAN_6987F and MXAN_6987R were used to amplify a 1,116-bp fragment of MXAN_6987 (Table S1). The amplicon was cloned into pCR2.1-TOPO (K4500-01; Invitrogen) to generate pHFS02 (Table 1). Plasmid pHFS02 was then transformed into electrocompetent DK1622 cells under Km selection. Plasmid pHFS02 was also transformed into strain LS4120 to generate the triple mutant LS4122.

Primers 5372UTR-F and 5372UTR-R were used to amplify the untranslated region of MXAN_5372, and tdT-F and tdT-R were used for tdTomato amplification (Table S1). The two DNA fragments were fused by overlapping PCR. The fused PCR product was digested with KpnI and BglII and then cloned into the same sites of pMR3487 to create pHFS03. This plasmid was transformed into electrocompetent DK1622, LS2442, LS3600, and SW2808 cells with Tc selection to generate the corresponding MXAN_5372 UTR::tdTomato strains: LS4134, LS4136, LS4138, and LS4140.

Sporulation and germination assay.

M. xanthus DK1622, LS4122, and LS4120 were grown in CYE broth to a density of 5 × 10⁸ cells · ml⁻¹. The cells were resuspended to 5 × 10⁹ cells · ml⁻¹ in deionized water, and 20-μl aliquots of cell suspension were spotted onto TPM agar plates and incubated at 32°C for 5 days. The cells were harvested and resuspended in 100 μl TPM buffer, incubated at 50°C for 2 h, and then subjected to sonication for 30 s to kill any remaining vegetative cells. The spores were then counted directly in a Petroff-Hauser counting chamber and plated on CYE agar plates using CYE soft agar (0.7% agar) overlays. The numbers of germinating spores were determined by counting colonies after 5 days at 32°C. The sporulation efficiency was evaluated as the percentage of the spores relative to that of direct DK1622 WT counts. The germination efficiency was given as the fraction of the viable spores relative to the direct count of each M. xanthus strain.

The production of spores for the mature spore stress tolerance assays was in accordance with the same general protocol as outlined above. To test the heat tolerance of 5-day-old spores, sonicated spores were incubated at 60°C or 65°C. The samples were removed at various time points over a 3-h period and plated on CYE using CYE soft agar overlays. The UV tolerance of spores was tested by exposing spores to 1,000 μW/cm² for up to 2.5 min while periodically removing samples for plating. After plating, the spores were incubated at 32°C for 5 days to allow germination and outgrowth.

Determination of rod and spore numbers.

M. xanthus DK1622, LS4120, and LS4122 strains were grown in CYE and then spotted onto TPM agar plates as described above. Every 24 h, the cells were scraped off the TPM agar plates and resuspended in 2% glutaraldehyde. The fruiting bodies were disrupted by a brief sonication. The numbers of rods and spores in each sample were then directly counted using a Petroff-Hauser counting chamber.

Lipid body quantification by Nile red staining.

Lipid bodies were observed with the lipophilic dye Nile red. A Nile red (Sigma-Aldrich) 0.5-mg/ml stock solution was prepared in 95% ethanol and kept in the dark. The stock solution was diluted to 1.25 μg · ml⁻¹ with water for lipid body staining (10, 11). M. xanthus cells were prepared for development on TPM agar as described above. At various time points, the cells were stained by adding Nile red directly to the top of the cell spot; this was followed by an incubation for 1 h at 32°C in the dark. The stained cells were imaged with a fluorescence microscope (DM5500B; Leica Microsystems). Digital images were obtained using a QICAM FAST 1394 camera (QImaging). The images were edited by using SimplePCI (Hamamatsu). The total fluorescence intensity of each cell or spore was calculated with Fiji (54). The experiment was repeated three times with at least 30 cells each to determine the mean fluorescence intensity.

FAD assay.

Flavin adenine dinucleotide (FAD) was quantified using a coupled enzyme assay (MAK035; Sigma-Aldrich). M. xanthus cells were grown to 5 × 10⁸ cells · ml⁻¹ in 1 liter CYE broth for approximately 24 h at 32°C. Then, the cells were centrifuged at 10,000 × g for 10 min and resuspended in fresh CYE to 5 × 10⁹ cells · ml⁻¹. Eight milliliters of this suspension was spread onto each of six 500-ml trays of TPM agar to induce development. After incubating at 32°C for 24 h, the cells were harvested and resuspended in 0.1 M potassium phosphate buffer (pH 6.4) and sonicated three times for 30 s each. Cell debris was removed by ultracentrifugation at 40,000 × g for 30 min. The bicinchoninic acid (BCA) protein assay reagent (23225X; Pierce) was used to determine the total soluble protein of different samples. Sample absorbance was measured at 570 nm every minute over the course of 30 min using a Synergy 2 plate reader (BioTek). The FAD concentration in each sample was normalized to the protein concentration in each sample separately.

tdTomato expression pattern analysis.

The expression of tdTomato (excitation/emission, ∼554/581 nm) was determined after starvation on TPM agar. Cells and spores were harvested from TPM agar plates at different time points and then dispersed and diluted for imaging. Images were obtained using a Leica Microsystems fluorescence microscope equipped with a QICAM FAST 1394 camera. The images were edited by using SimplePCI, and the fluorescence intensity of each cell or spore was calculated with Fiji and is expressed in arbitrary units (AU). The mean fluorescence intensity is derived from measurements of 30 cells or spores.

Electron microscopy.

M. xanthus DK1622, LS4122, and DK851 were grown in CYE broth to a density of 5 × 10⁸ cells · ml⁻¹. The cells were resuspended to 5 × 10⁹ cells · ml⁻¹ in deionized water, and 20 μl of the cell suspension was spotted onto TPM agar plates and incubated at 32°C for 48 h or 144 h. After incubating, the fruiting bodies were harvested and then lysed by sonication to release spores, as described above. The spores were rinsed once in TPM and then fixed overnight in 2% (vol/vol) glutaraldehyde in 0.1 M cacodylate buffer. The samples were washed three times in 0.1 M cacodylate buffer before postfixing in 2% OsO₄ for 1 h. After the samples were washed five times in H₂O, they were stained for 1 h in 0.5% uranyl acetate. The samples were washed in water three more times and then dehydrated using a graded series of ethanol (30, 50, 75, and 96% and twice with 100%) for 15 min each. The samples were washed in 100% propylene oxide twice and then embedded using Spurr resin at propylene oxide ratios (vol/vol) of 25:75, 50:50, and 75:25 for 6 to 8 h each and finally in 100% Spurr for 24 h. After curing for 6 h at 70°C, the samples were mounted, sectioned, and viewed with a JEOL JEM-1011 transmission electron microscope (JEOL, Inc., Peabody, MA) at 80 kV. This protocol was adapted from those in references 11 and 55.

The spore coat thickness was calculated from electron micrographs. The spore coat thickness was measured manually in a minimum of three places, always including the thickest and thinnest portions of the spore coat. The thickness in μm was calculated on the basis of the magnification and scale of the image. These values were then averaged to provide the average thickness of each spore coat. The thicknesses of at least 20 spore coats were averaged together to calculate the average spore coat thickness of each strain.