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Infection and Invasion of Roots by Symbiotic, Nitrogen-Fixing Rhizobia during Nodulation of Temperate Legumes

Department of Molecular and Cell Biology, University of Connecticut, Storrs, Connecticut 06269

SUMMARY

INTRODUCTION

Topics Covered in This Review

Brief Outline of the Infection Process in Plants That Form Indeterminate Nodules

GROWTH OF ROOT HAIRS

Cytology and Development of Root Hairs

Roles of Actin and Microtubule Cytoskeleton in Tip-Growing Cells

Roles of Calcium in Tip-Growing Cells

Roles of Other Proteins in Tip-Growing Cells

NADPH oxidase.

Small GTPases.

INITIATION OF INFECTION

Adhesion of Rhizobia to Root Hairs

Root Hair Deformation and Curling

Is There a Nod Factor-Mediated Signal That Is Specifically Required for the Initiation of Infection Threads?

Degradation of Cell Wall Associated with Infection Thread Initiation

Questions about Initiation of Infection Threads

EXTENSION OF INFECTION THREADS THROUGH ROOT HAIRS

Plant Contributions to Infection Threads

Bacterial Contributions to Infection Threads

Growth Dynamics of Bacterial Populations inside Infection Threads

INFECTION THREAD INVASION OF THE DEVELOPING NODULE

Early Steps of Infection Thread Invasion: Activation of the Pericycle and Root Cortex

Later Steps: Growth and Architecture of Infection Thread Networks in Young Nodules

PLANT CONTROL OF INFECTION

Plant Defense Responses following Infection Thread Formation

Ethylene and Control of Infection and Nodulation

ECOLOGICAL QUESTIONS ABOUT RHIZOBIAL INFECTION

OUTLOOK

ACKNOWLEDGMENTS

REFERENCES

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INTRODUCTION

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Prior to agriculture, biological fixation of atmospheric nitrogen is estimated to have accounted for 90% of the 100 to 140 Tg of nitrogen (1 Tg = 10¹² g [10⁶ metric tons]) fixed annually in terrestrial environments. The remaining 10% was fixed abiotically, primarily by lightning. Now human activity, especially the generation of ammonium compounds for agricultural fertilizers, fossil fuel consumption, and increased planting of legumes, contributes an estimated 140 Tg of additional fixed nitrogen each year (179). Biological nitrogen fixation is catalyzed by prokaryotes only, so far as is known. The group of prokaryotes that do this is large and diverse and contains both eubacteria and archaea (186, 193). Nitrogenase, the enzyme complex responsible for nitrogen reduction, is irreversibly inactivated by oxygen; therefore, this process requires conditions that are anoxic or nearly anoxic. In oxic environments nitrogenase is protected from inactivation by being sequestered in differentiated cells with morphological and biochemical characteristics that limit exposure of nitrogenase to oxygen. In some plants, root nodules develop to house nitrogen-fixing bacteria in a microaerobic environment. This process, a type of symbiotic nitrogen fixation, is, for the most part, restricted to a limited number of bacterial groups, including the genera Rhizobium, Mesorhizobium, Sinorhizobium, Bradyrhizobium, and Azorhizobium (collectively referred to in this review as rhizobia) and Frankia. All but the last of these are from the -proteobacterial Rhizobiaceae family and induce nodules on plants from the Leguminosae family. Frankia is a filamentous gram-positive actinomycete that induces nodules on a variety of woody plants from the Betulaceae, Casuarinaceae, Myricaceae, Elaegnaceae, Rhamnaceae, Rosaceae, Coriariaceae, and Datisticaceae families (10, 11). Rhizobia carry most of the genes specifically required for nodulation either on large (500-kbp to 1.5-Mbp) plasmids or on symbiosis islands (4, 58, 85, 86). Interestingly, it has been recently discovered that bacteria from outside the Rhizobiaceae can induce nodules on legumes. For example, a strain of Methylobacterium, an -proteobacterium, can nodulate Crotalaria, and ß-proteobacteria related to Burkholderia can nodulate Machaerium lunatum and Aspalathus carnosa (110, 159). Apparently these species have acquired, by horizontal gene transfer, plasmids or islands that contain many of the same genes used by typical Rhizobiaceae to induce nodule formation and catalyze nitrogen fixation (110, 159).

Nodules induced by rhizobia are of two general kinds, determinate and indeterminate. These differ in a number of respects, one of the most important being that indeterminate nodules are elongated and have a persistent meristem that continually gives rise to new nodule cells that are subsequently infected by rhizobia residing in the nodule. These newly infected cells, and the bacteria inside them, develop further and form new nodule tissue that actively fixes nitrogen. This process results in a gradient of developmental stages, from the young meristem at the nodule tip to the older senescent tissue near the root (Fig. 1).

View larger version (50K): [in this window] [in a new window]   FIG. 1. Overview of the nodulation process in plants that form indeterminate nodules. (A) One form of Nod factor synthesized by S. meliloti. The upper arrow indicates the acetyl group added by NodL, and the lower arrow indicates the lipid moiety, the length and degree of saturation of which is modified by NodF and NodE. (B) Diagrammatic cross section of a root, showing gradients of an activating factor at protoxylem poles (blue) and an inhibitor at protophloem poles (red). Together such gradients may determine which root cells can become activated in response to infecting rhizobia. Nodules are typically formed next to the protoxylem poles, which are at the ends of the Y-shaped structure depicted in the center the diagram, rather than above the protophloem poles, which are depicted as ovals. (C) An epidermal cell and two underlying outer cortical cells. The epidermal cell has a nucleus positioned across from the place where a new root hair will form. (D) Root hair initiation in the epidermal cell. (E) Binding of a rhizobial cell to a type I root hair and activation of the underlying cortical cells in response to Nod factor. (F) Continued growth of the root hair, shown as stage II. (G) Curling of the stage II root hair under the influence of Nod factor and growth of a rhizobial microcolony in the curl. The underlying cortical cells have become polarized, and cytoplasmic bridges (PITs) have formed and are shown in grey. (H) Infection thread initiation. (I) Growth of the infection thread down the root hair. The nucleus moves down the root hair in front of the thread. (J) Fusion of the infection thread with the epidermal cell wall and growth of rhizobia into the intracellular space between the epidermal cell and the underlying cortical cell. (K) Growth of the infection thread through PITs in the outer cortical cells. (L) Enlarged view of the root hair shown in panel I. The curl has been unrolled to show that topologically, the bacteria in the infection thread are still outside the root hair. The plant cell wall and plant cell membrane are shown as black and dashed lines, respectively. Microtubules are located between the nucleus and the infection thread tip (blue). Actin cables are depicted as orange strands. These are likely to be found where indicated in the diagram because cytoplasmic streaming is seen in these areas during the progression of infection threads down root hairs. Bacteria are topologically outside the root until they later bud from the tip of the thread and enter nodule cells as membrane-enclosed bacteria. (M) Diagram showing root tissues and a young nodule not yet emerged from the root. The derivation of nodule tissues from root tissues is indicated. (N) A longitudinal section of 10-day-old alfalfa nodule. The nodule was infected with GFP-expressing S. meliloti, and the infection thread network can be seen behind the meristem region of the nodule. The initial infection site that gave rise to the bacteria in the nodule can be seen on the nodule periphery at the left. Propidium iodide (red) was used to counterstain the plant tissue. The root from which the nodule emerged is seen in cross-section at the right.

Determinate nodules lack a persistent meristem, are usually round, and do not display an obvious developmental gradient as do indeterminate nodules. Legumes that form determinate nodules are typically tropical in origin and include Glycine max (soybean), Vicia faba (bean), and Lotus japonicus. L. japonicus, like M. truncatula, has genetic characteristics that make it particularly suitable as a model to study the formation of determinate-type nodules (71).

Other responses of rhizobia upon encountering a host root undoubtedly involve changes in the expression of genes other than those involved in Nod factor synthesis. Such genes are likely to be important for rhizobia to compete effectively with other organisms for access to growth substrates emanating from the host root, to adhere to the root surface, and to become resistant to toxic substances such as phytoalexins secreted by the root.

Early during symbiosis, rhizobia must get from the root surface to the inner root tissue where they will populate cells in the incipient nodule. To do this, they grow and divide inside a tubule called an infection thread. Infection thread formation is most often initiated when rhizobia become trapped between two root hair cell walls. This usually occurs when a deformed root hair forms a sharp bend or curl, and bacteria bound to the root hair become trapped between appressed cell walls (26). Invagination of plant cell wall in the curl, or degradation of the wall and invagination of the cell membrane, followed by tip growth of the invagination results in the initiation of an infection thread that grows down the inside of the root hair and into the body of the epidermal cell. Rhizobia inside the thread grow and divide, thereby keeping the tubule filled with bacteria. If the infection thread exits the epidermal cell, it does so by fusing with the distal cell wall, and bacteria enter the intercellular space between the epidermal cell and the underlying cell layer. Invagination and tip growth, similar to those seen at the beginning of infection thread growth, occur in the underlying cell, and a thread filled with bacteria is propagated further toward the root interior (170, 173, 175). Branching of the thread as it grows through the root and enters the nodule primordium increases the number of sites from which bacteria can exit the thread and enter nodule cells, ensuring that a sufficient number of nodule cells are colonized. Bacteria inside the infection thread eventually exit it and enter nodule cells. Once inside nodule cells, the bacteria continue to differentiate and synthesize proteins required for nitrogen fixation and for the maintenance of the mutualistic partnership. The processes described above are outlined in Fig. 1.

In the sections that follow, the growth of root hairs, their curling under the influence of Nod factor, initiation and growth of infection threads, growth of infection thread networks through the root cortex and in nodules, and plant control of infection are covered in detail. Some related areas are not covered, but the reader is provided with citations to current reviews on those topics.

GROWTH OF ROOT HAIRS

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Root hairs and pollen tubes are the best-studied plant cells that elongate through the process of tip growth (28, 75, 156). Infection threads develop from growing root hairs, are thought to be tip-growing structures, and therefore most likely elongate by using at least some of the machinery that was supporting root hair growth before infection took place. Thus, understanding the processes that contribute to tip growth in root hairs and pollen tubes should shed much needed light on processes involved in infection thread growth.

View larger version (57K): [in this window] [in a new window]   FIG. 2. Root hair morphology on uninoculated and inoculated roots. (A to C) Typical root hairs from zones I, II, and III, respectively, of an uninoculated alfalfa plant. (D) Diagram of an alfalfa seedling, showing the locations of root hair zones I, II, and III. (E to G) Photographs showing how root hair responses to S. meliloti can vary along the length of a single root. The three images were taken from a single inoculated seedling at the locations indicated on the central diagram.

Vesicles that fuse at the tip of root hairs are derived from Golgi bodies that are a short distance behind the growing tip. The vesicles deliver membrane and cell wall components that are incorporated into the plasma membrane, cell wall, and extracellular matrix. During tip growth, actin-dependent cytoplasm streaming brings vesicles and other organelles to the apical region of the cell. The cytoplasm typically moves toward the growing tip along the outside of the cell and then moves back toward basal regions through the center of the cell. This pattern, most obvious in pollen tubes, is referred to as reverse fountain streaming (75, 83). The region immediately adjacent to the tip does not exhibit cytoplasmic streaming and is free of organelles that can be resolved by typical light microscopy. This region, termed the clear zone, contains the vesicles that fuse with the tip and deliver the material required for growth (reviewed in reference 107). Because the vesicles are delivered to the base of the clear zone and are consumed at its apex, it has been suggested that diffusion alone may suffice to transport the vesicles from the base of the clear zone to their site of fusion near the tip of the root hair (97, 107).

The role of microtubules in tip growth is less clear than is the role of actin. This is because microtubule-stabilizing and -destabilizing drugs can cause dramatic growth distortions in root hairs and pollen tubes but often do not stop tip growth (13, 102, 150). Work with Arabidopsis root hairs led Bibikova et al. to the conclusion that actin filaments were required for tip growth and microtubules were required for the proper directional control of the extending tips (13). The effects of microtubule-stabilizing and -destabilizing drugs on the morphology of growing M. truncatula root hairs have shown that microtubules are involved in maintaining the normal structure of the subapical cytoplasmic dense region and are involved in maintaining a normal distance between the nucleus and the growing tip of the root hair (150). Microtubules may also organize actin filaments during root hair growth (102, 162). This has been suggested because when both the actin and microtubule networks are depolymerized, and then actin filaments, but not microtubules, are allowed to reform, cytoplasmic streaming is abnormal (162).

In spite of the accumulating data correlating the Ca²⁺ gradient with tip growth, the actual roles played by Ca²⁺ in tip growth are still somewhat obscure. Suggested roles for Ca²⁺ include influencing cell wall strength and extensibility, controlling actin dynamics, and controlling vesicle fusion at the growing tip (28, 75, 138). Of course, these roles need not be mutually exclusive.

Small GTPases. Rac/Rho-like small GTPases (called ROPs, for Rho of plants) are found in plants and have been implicated in a variety of functions, including signal transduction, control of actin dynamics, and control of tip growth in root hairs and pollen tubes (59, 70, 92, 192). These proteins have been localized to growing tips of root hairs and pollen tubes. Interference with normal activity of some of the family members affects polarized growth of both of these cell types. For example, in Arabidopsis, which has 11 ROPs (156), expression of constitutively active ROP4 or ROP6 in Arabidopsis causes depolarized growth of root hair tips resulting in swollen, distorted tips; the distortions were correlated with delocalized calcium gradients in the tips (108). Similar experiments have implicated ROP2 and ROP7 in control of tip growth of Arabidopsis root hairs (84). Thus, the evidence suggests that at least some of the ROPs involved in control of tip growth may act by affecting actin dynamics and/or calcium gradient formation in the apical region of growing of root hairs and pollen tubes.

INITIATION OF INFECTION

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For the purpose of this review, initiation of infection is considered to encompass the steps from bacterial adhesion to root tips through the beginning of infection thread growth. Many of these infection events are mediated by Nod factor. Recent papers have covered Nod factor structures, the synthesis of Nod factors, Nod factor signal transduction, and the relationship between host range and Nod factor structure. The reader is directed to previously published reviews (20, 38, 63, 97, 146, 165, 181, 183) for a comprehensive discussion of these topics.

Following weak binding, a tight binding step that is mediated by the bacterial synthesis of cellulose fibrils is initiated (153, 154). The synthesis of these fibrils was shown to be required for R. leguminosarum to form biofilm-like caps on the tips of pea root hairs. Mutants that did not form the fibrils did not form caps, but they were able form nitrogen-fixing nodules, which indicated that capping and cellulose-mediated tip binding are not absolutely required for a successful symbiosis to occur (154). However, binding and capping may be needed for rhizobia to effectively colonize root hairs under natural conditions where competition for access to root surfaces and grazing pressure by eukaryotes are likely to be intense.

Host lectins have also been shown to play roles in rhizobial adhesion to plants that form determinate and indeterminate nodules. These lectins localize to root hair tips and are thought to help convey host-symbiont specificity by binding simultaneously to the plant cell wall and to saccharide moieties on the surfaces of compatible bacteria (41, 43, 79). A series of experiments in which a variety of transgenic plants expressed lectins from other species of legumes has shown that the presence of heterologous lectins often allows transgenic plants to respond to symbionts that are usually noncompatible, provided that the heterologous lectin can bind to the noncompatible bacteria and provided that the noncompatible bacteria make the proper Nod factor (42, 79, 171, 172). For example, transgenic alfalfa that expressed pea lectin formed nodules and infection threads when inoculated with low numbers of R. leguminosarum bv. viciae that synthesized Sinorhizobium meliloti Nod factors. Control plants and transgenic plants that expressed mutant pea lectin did not form nodules or infection threads when inoculated with low numbers of the same strain (171). Results such as these suggest that cell-cell contact and specific binding of compatible bacteria to root tips are important for infection and infection thread formation, because they result in the exposure of infectible root hair tips to the proper symbiont and hence to a high localized concentration of the Nod factors needed to trigger root hair curling and infection thread formation (79, 171). A direct test of the importance of host lectins in promoting infection and invasion would be very interesting. Such a test may be difficult, however, because down regulation of lectin expression causes severe developmental defects in alfalfa and may (or may not) also do so in other legumes (19).

Nod factor induced deformation of zone II root hairs begins with root hair tips swelling isodiametrically; this is followed by the establishment of a new growing tip that resembles highly polarized, actively growing tips of zone I root hairs (40, 72, 106, 149). Thus, purified Nod factor can induce new tip growth in zone II cells. It is not yet clear why vesicle deposition becomes temporarily isodiametric following exposure to Nod factor. Isodiametric deposition may come about because of the disruption of the cytoskeleton. An alternative explanation, set forward by Sieberer and Emons, is that zone II root hairs normally no longer have a tip-directed vesicle delivery system and, upon Nod factor addition, Golgi vesicles are incorporated randomly into the tip region until a new tip-focused delivery system is reestablished (149).

It is interesting to consider why zone I and zone III cells do not deform in response to the addition of purified Nod factor. Zone III cells may be unable to deform because they have a secondary cell wall or because they no longer have the machinery in place to catalyze tip growth. Zone I cells may not respond to Nod factor by deforming because they are already highly polarized and actively growing (97). It has been pointed out that root hairs in zones I and III change in other ways after exposure to Nod factor and are thus responsive to its presence. For example, zone I hairs exhibit depolarization, Ca²⁺ influx, and Ca²⁺ spiking (47, 48, 53, 146, 183), and the subapical region vetch root hairs in zones I, II, and III exhibit a characteristic increase in fine bundles of actin following addition of Nod factor (39).

When added to the external medium, purified compatible Nod factors are sufficient to cause root hair deformation and branching, but they are not sufficient to cause the formation of tightly curled root hairs (shepherd's crooks) that are usually the sites of bacterial entry into plants. It had been hypothesized that the reason for this is that a localized source of Nod factor is needed to continually redirect the off-axis tip growth needed to form a tight curl (49, 135, 168). This idea has intuitive appeal. Recent experiments in which purified Nod factor from S. meliloti was applied in a highly localized fashion to root hairs of M. truncatula have shown that a point source of Nod factor can cause root hairs to grow into structures resembling shepherd's crooks (50).

Following inoculation with compatible bacteria, root hairs on a single plant can show a wide range of deformation morphologies. Hairs in some regions of the root show no deformation at all, other regions have wavy root hairs with swellings, and other regions are populated with tightly curled root hairs (shepherd's crooks) that are able to support the development of infection threads (Fig. 2) (26). These various morphologies are often seen in a single plant whose root hairs went through all phases of development in the presence Nod factor-secreting bacteria. Assuming that Nod factor was continually present, these observations suggest that root hair responsiveness toward Nod factor changes as the root grows. The responsiveness of root hairs to deform in the presence of Nod factor can be modulated by plant hormones such as ethylene, which inhibits Nod factor signal transduction and can influence the degree of root hair deformation and the frequency of productive infections (119) (see below). Thus, changes in ethylene levels, in ethylene signal transduction, or in other hormone signaling systems during root growth may explain the observed variability in root hair responsiveness to Nod factor.

It was shown by Cardenas et al. that Nod factor causes the fragmentation of the actin microfilament network upon addition to bean root hairs (27). The actin bundles normally seen in bean root hairs injected with fluorescent phalloidin disappeared within 5 min after the application of Nod factor, and an area of diffuse fluorescent staining concomitantly appeared in the apical region of the root hair. Because phalloidin binds to filamentous actin, this diffuse staining presumably represented filamentous, fragmented microfilaments. Actin bundles reappeared within an hour of Nod factor addition, indicating that cells were able to at least partially recover a microfilament network even when Nod factor was present (27).

Membrane depolarization, Ca²⁺ influx at the root hair tip, and Ca²⁺ spiking in the perinuclear region occur within seconds to minutes upon addition of compatible Nod factors to growing legume root hairs. It has been hypothesized that the influx of calcium associated with Nod factor addition may be related to the fragmentation of the actin network at the root hair tip, as is the case with pollen tubes (90).

Work done by de Ruijter et al. with vetch leads to a different picture of how the actin network responds to the addition of Nod factor. In this case, quantitation of actin bundles in root hairs fixed and then stained with phalloidin 3 to 15 min after addition of Nod factor revealed that the root hairs responded to Nod factor by increasing the number of fine actin filaments in the subapical region. The authors suggested that such an increase could have come about by unbundling actin cables or by polymerization of actin filaments (39). The conflicting observations of Cardenas et al. and de Ruijter et al. have yet to be reconciled. Species differences or the fact that one study employed injected phalloidin for visualization and the other employed phalloidin staining after fixation may explain the different responses observed by the two groups. Given the critical role of the actin network in tip growth, it will be important to understand exactly how this network responds to Nod factor and how that response relates to root hair deformation and curling.

In addition to changes in the actin component of the root hair cytoskeleton, the microtubule component also changes following exposure to compatible rhizobia. A detailed study, employing antibodies against tubulin, showed that arrays of microtubules changed after M. truncatula root hairs were exposed to wild-type S. meliloti. Initially microtubules were arranged in the root hairs in helical cortical arrays. Following exposure to S. meliloti, the microtubules in root hair cells rearranged to form endoplasmic networks around nuclei and cortical networks that were no longer helical but were more closely aligned with long axes of the root hairs. Nuclei and their associated microtubules then moved into the tip region of the root hairs, and the microtubule array became positioned between the tip of the root hair and the nucleus. As curling commenced, the endoplasmic microtubule arrays became associated with the center of the curl. If the curl developed an infection thread, the microtubular array became disconnected from the root hair tip and instead connected the tip of the infection thread to the nucleus (161). This arrangement likely remains as nuclei advance down root hairs in front of infection threads.

It is too soon to integrate information about root hair growth and the events of Nod factor-induced deformation into a detailed model that explains the molecular and cellular events leading to the formation of the tightly curled root hairs that support rhizobial infection. However, any such model must take into account how rearrangements of the actin and microtubule networks lead to deposition of cell material at off-axis sites that are away from the ring of deposition that is normally near the root hair tip (145). Given that (i) actin directs cell material to the tip of the growing root hair, (ii) actin may fragment and reassemble after the addition of Nod factor, (iii) the microtubule network rearranges to become centered on the growing curl, and (iv) the microtubule network is involved in stabilizing the growth site at the root hair tip, it is reasonable to propose that redirection of tip growth may require the breakup of the preexisting actin network at the root hair tip, followed by its reassembly at on off-center site that is stabilized, or selected, by the microtubule network, which is itself is rearranged during the early steps of rhizobial infection. The ultimate signal for these spatial reorganizations would be bacteria bound to the root hair tip, acting as a point source of Nod factor and perhaps as a point source of other bioactive signals as well (49, 50, 135, 168).

Ardourel et al. (2) proposed a model in which root hairs have at least two receptors or signal transduction pathways devoted to Nod factor perception. One is less stringent and, when activated by Nod factor or Nod factor with an altered nonreducing end, causes root hair deformation and cortical cell division. The other is more stringent and is activated by wild-type Nod factor but not by Nod factor synthesized by the nodF nodL strain. This more stringent pathway is required for shepherd's crook formation and infection thread initiation, and it was postulated to inhibit the deformations and cortical cell divisions induced by the first pathway. Thus, Nod factors made by nodF nodL strains result in deformation and cortical cell division but are not sufficient for infection thread formation or for down regulation of deformation and induced cortical cell division.

In a related study, Walker and Downie inoculated a nodFEMNTLO deletion mutant of R. leguminosarum bv. viciae onto vetch seedlings (182). The mutant synthesized Nod factor molecules that contained no host-specific decorations. Microscopic observation of roots showed that inoculated plants contained overly deformed root hairs and occasionally formed shepherd's crooks that often contained large aggregations of bacteria, but they failed to initiate infection threads from such crooks. Interestingly, this was similar to the phenotype of the nodF nodL double mutants of S. meliloti when they were inoculated onto M. truncatula. Note that this was different than the response seen with the nodF nodL mutant on alfalfa, where shepherd's crooks were not initiated. The phenotype seen when the nodFEMNTLO stain was inoculated onto vetch is particularly intriguing result because it again shows that Nod factor decorations are not needed for root hair deformation but are required for the initiation of infection threads.

Walker and Downie (182) showed that overexpression of nodO rescued nodFEMNTLO phenotypes to some extent. With nodO overexpression the mutant no longer accumulated in large masses, some root hairs initiated infection threads, and some nodules formed. NodO protein is known to be secreted from bacterial cells, has been shown to form pores in membranes, and can be a host range determinant (180). It may be that NodO rescued the nodFEMNTLO phenotype by allowing ion flow across a root hair membrane and thus amplifying a weaker-than-normal signal transmitted by the stripped-down version of Nod factor.

Recently, strong candidates for the stringent Nod factor receptor required for infection thread initiation in M. truncatula have been identified by Limpens et al. (101). Putative genes (LYK3 and LYK4) for this receptor encode two similar histidine kinases with extracellular LysM domains. The LysM domains are most similar to LysM domains of a Volvox carteri chitinase gene. This similarity is interesting given that the undecorated backbone of Nod factor is a small molecule of chitin. Using RNA interference, the Limpens et al. down regulated the expression of LYK3 and LYK4. The root hairs of transgenic LYK3 and LYK4 knockdown mutants had aberrant infection threads when inoculated with S. meliloti nodFE mutants, which synthesize Nod factors with an alternative lipid (vaccenic acid C_18:1 11) at the nonreducing end. The structure of the aberrant threads was very similar of those of threads induced by the S. meliloti nodF nodL double mutant discussed above (2). On wild-type roots the S. meliloti nodFE mutant was able to induce infections and infection threads that were indistinguishable from those induced by wild-type S. meliloti. Wild-type S. meliloti also induced aberrant threads on the LYK3 mutant, although not at the same frequency as the nodFE mutant, indicating that the LYK3 knockdown phenotypes were not induced only in response to the nodFE mutant. In addition, 20% of the nodFE microcolonies in shepherd's crooks on LYK3 knockdown roots initiated infection threads, whereas 80% of the microcolonies initiated infection threads on wild-type roots. Together, the data presented by Limpens et al. are compatible with the hypothesis that the nonreducing end of S. meliloti Nod factor interacts with LYK3 or LYK4 and that the signal resulting from such an interaction is required to initiate infection threads and for proper polar growth of those infection threads.

Two L. japonicus genes encoding histidine kinases with LysM domains, NFR1 (Nod factor receptor) and NFR5, were recently shown to be required for the earliest plant responses to Nod factor and the microsymbiont of L. japonicus, Mesorhizobium loti (103, 129). The genetic and physiological characteristics of the NFR mutants are consistent with their encoding a Nod factor receptor needed for the first steps of symbiosis. The NFR1 and NFR3 proteins are very similar to the LYK3 and LYK4 proteins of M. truncatula, but the phenotypes associated with the NFR mutants are much more extreme. At this point it is not clear if the phenotypic differences between the NFR mutants and the LYK mutants are because the proteins, although similar, are required at different steps of the symbiosis, because the LYK knockdown mutants retained some residual activity that allowed steps such as root hair deformation to occur, or because the signal transduction pathways of the two plants may be different in terms of redundancies or other key properties.

The question of which partner, plant or bacterium, is responsible for the degradation remains open. Rhizobial bacteria have enzymes that are capable of degrading cellulose and other plant cell wall polysaccharides (82, 105, 109, 127, 194). Studies by Mateos et al., employing electron microscopy, have shown that R. leguminosarum bv. trifolii and S. meliloti induce the formation of pits on root surfaces of their respective hosts (104). These pits are the same size and shape as the bacteria and are found directly under bacteria bound to roots, indicating that if the bacteria are producing the enzymes responsible, the enzymes are bound to the bacterial surface and are not secreted. It should be noted that electron micrographs depicting the degradation associated specifically with infection thread initiation show that degradation, while localized, is more widespread and diffuse than that shown by Mateos et al. Plants clearly have the capability to degrade, or otherwise alter, their cell walls. For example, cell walls are weakened or degraded during root hair initiation, fruit ripening, pollen tube transit down the pistil, and leaf abscission (21, 34, 35, 137). Observations that cell wall-degrading enzyme activities, induced by compatible bacteria, do not occur in the presence of nitrate are easiest to explain if the host has at least some control over degradation in response to rhizobial infection (52, 105, 173).

It was recently shown that an alfalfa polygalacturonase gene (MsPG3) was induced specifically in roots in response to S. meliloti (111). The gene was induced as early as 1 day after spot inoculation, and high levels of expression of this gene were seen in the nodule meristem, infection zone, and interzone II-III of nodules. Those authors did not determine whether the gene was induced in root hairs following inoculation. Given that cell wall degradation likely occurs before infection thread initiation, it would be interesting to know if MsPG3 is induced in root hairs following inoculation with S. meliloti.

EXTENSION OF INFECTION THREADS THROUGH ROOT HAIRS

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Given that infection threads initiate only in growing root hairs, it is likely that some of the machinery required for root hair extension is also required for infection thread extension. During growth of infection threads down root hairs, there is an association between the extending infection thread tip and the root hair nucleus, much as there is a connection between the root hair tip and the root hair nucleus during growth of root hairs.

During growth of infection threads down root hairs, the nucleus is connected to the extending infection thread tip by a thick and actively streaming column of cytoplasm (Fig. 3A). While the roles of actin and microtubules in root hair growth and associated cytoplasmic streaming have been investigated, their roles in infection thread growth remain uninvestigated. Because of the active cytoplasmic streaming in the tip region of extending threads, plant cytoskeleton is likely to have an important role in growth of infection threads. In alfalfa, the large amount of active cytoplasm in the tip region and immature infection thread cell wall makes discerning the tips of growing threads difficult. In some cases only thin cytoplasmic strands can be seen connected to threads with well-defined tips (Fig. 3B). In alfalfa, such threads with well-defined tips have invariably stopped growing through root hairs (C. Arango and D. J. Gage, unpublished data).

View larger version (123K): [in this window] [in a new window]   FIG. 3. Infection threads in alfalfa root hairs. (A) A growing infection thread, with an active column of cytoplasm between the nucleus (arrow) and the tip region of the thread (arrowhead). The actual tip of the thread cannot be discerned because it is obscured by the cytoplasmic column or because it has not yet been enclosed with thick, mature cell wall. The inset shows the nucleus and its associated column of active cytoplasm. (B) An infection thread that has terminated growth. Note that the tip is easily discerned and the tip region has only very thin cytoplasmic streams connected to it (inset).

Studies of the hcl (hair curling) mutant of M. truncatula have shown that its root hairs deform and branch after inoculation with S. meliloti but that subsequent curling and infection steps do not occur. Shepherd's crook formation, infection thread initiation, infection thread extension, and the concomitant reorganization of microtubule arrays were not present following inoculation, suggesting that morphological changes of root hairs during infection and reorganization of microtubule arrays are directly or indirectly linked (30). Those authors hypothesize that hcl mutants are deficient in establishing or responding to the organizational signals that emanate from S. meliloti microcolonies and that are normally needed for shepherd's crook formation, infection thread initiation, reorganization of microtubule arrays in root hairs, and polarization of root cortical cells (30).

Alfalfa and M. truncatula plants fail to nodulate properly when inoculated with S. meliloti Rm1021 exo mutants that are unable to produce succinoglycan, an exopolysaccharide consisting of repeating acidic octcasaccharide subunits (54, 95). Nodules induced by such strains are small, lack a persistent meristem, contain few or no bacteroids, and cannot fix nitrogen (54, 95). The nodules are empty, because S. meliloti exo mutants are inefficient at initiating infection threads. Those that do initiate are bloated and grossly misshapen and usually cannot extend beyond the basal part of the root hair (Fig. 4) (31, 121, 190). Infection threads also cannot initiate normally in vetch root hairs inoculated with R. leguminosarum bv. viciae exo mutants (175). In the S. meliloti-alfalfa symbiosis and other symbioses, the Fix^– phenotypes of exo mutants can be reversed by the addition to roots of small amounts of purified, low-molecular-weight succinoglycan (6, 44, 65, 167). Fractionation studies of S. meliloti succinoglycan indicated that small quantities of the trimeric form of the repeating octasaccharide were sufficient for extracellular complementation of exo mutants. Two conclusions can be drawn from the fact that only small quantities of succinoglycan were sufficient for complementation of exo mutants: (i) that signaling is an important function of trimeric form of succinoglycan and (ii) that succinoglycan is unlikely to be a required structural component of the infection thread matrix (6, 44, 65, 167, 184).

View larger version (68K): [in this window] [in a new window]   FIG. 4. Infection threads induced by an exoF mutant of S. meliloti. (A) Two swollen and misshapen infection threads induced by a GFP-expressing exoF mutant of S. meliloti. Arrows point to sections of the threads that are particularly swollen. (B) Another example of an infection thread induced by the exoF mutant. For infection threads induced by wild-type S. meliloti on alfalfa, see Fig. 5.

K antigen, a capsular polysaccharide, is not synthesized by S. meliloti strain Rm1021 but is made by other S. meliloti strains. Analysis of strain Rm41, a strain that makes succinoglycan and K antigen, and of isogenic mutants that make only K antigen shows that this exopolysaccharide can also support infection thread initiation, infection thread extension, and nodule development (121).

The roles that succinoglycan, EPS II and K antigen might have in infection thread initiation and growth are not clear. The misshapen infection threads, overfilled with bacteria, that are often seen in the absence of succinoglycan (Fig. 4) suggest that the growth of the bacteria inside the threads is not matched to the growth of the tube (31, 121). This may be because the rate at which the tube is synthesized, or extends, is abnormally low, or its growth polarity may be altered. Pellock et al. suggested that succinoglycan may be involved in organizing the root hair cytoskeleton in such a way that tip growth of the infection thread is rapid and highly polarized (121). Others have shown that exopolysaccharides are able to modulate (down regulate) defense responses in plants (57, 114, 115, 174, 175). Thus, rhizobial mutants lacking exopolysaccharides may cause abnormal infection thread growth because the plant mounts an inappropriate defense response to the invading rhizobia.

Nod factor is required for the redirection of root hair growth that leads to shepherd's crook formation. If infection thread initiation and growth result from a continual redirection of root hair cell wall growth, then Nod factor may play an important role in these processes as well. The fact that mutations that alter the nonreducing end of Nod factor affect infection thread initiation and extension supports this idea (2, 182). That the LYK3 and LYK4 knockdown mutants are altered in infection thread initiation and extension may also indicate that Nod factor production is important for processes beyond root hair deformation (101). Finally, work showing that nod genes are expressed by bacteria in tissues of the nodule that contain actively growing infection threads supports the idea that infection thread propagation probably requires Nod factor-dependent targeting of material to infection thread tips in these cells as well (143, 144).

The use of S. meliloti tagged with green fluorescent protein (GFP) and similar proteins has rendered bacterial growth inside the infection threads particularly amenable to study (31, 60, 61, 121, 189). Upon mixing fluorescent and nonfluorescent S. meliloti, infection threads that contain both types of bacteria develop. Tagged bacteria often enter the infection thread and form alternating light and dark sectors with sharp boundaries between the sectors (Fig. 5) (61). These sectors apparently originate from the clonal expansion of founder cells which enter the thread very early. Recent work has confirmed these observations and extended them by showing that even though sectored infection threads are the most common type of mixed infection thread, other types of mixed threads can form, including ones in which strains appear to be randomly mixed (Fig. 5) (60). Observations of infections arising from bacterial strains tagged with different fluorescent proteins have led to insights into the infection and invasion process because they allow spatially distinct subpopulations of bacteria to be discerned. Growth patterns of the whole population in infection threads can be inferred from the growth and behavior of the observed subpopulations. For example, observation of threads populated with two S. meliloti strains that could be differentiated by fluorescence microscopy indicated that most of the bacterial cells in infection threads were not growing. Bacteria near the extending tips of threads were growing; the rest, near the back of the threads, appeared to be static (60, 61). The sizes of these bacterial growth zones were estimated to be about one-quarter to one-third of the length of a typical root hair (60).

View larger version (53K): [in this window] [in a new window]   FIG. 5. Examples of mixed infection threads following coinoculation of alfalfa with red- and green-fluorescent bacteria. (A) Infection threads containing only gfp-expressing or DsRed-expressing S. meliloti. (B) A sectored infection thread in which the mixed population gave rise to a series of sectors which increase in length along the thread. The tipmost sector advanced into the epidermal cell body, branched (top arrow), and penetrated the underlying cell (bottom arrow), leaving the distal sectors behind. (C) A root hair infected with dual infection threads. Each thread contains sectors of gfp-expressing and DsRed-expressing bacteria. (D) A jumbled type of mixed infection thread. gfp-expressing and DsRed-expressing cells appear randomly mixed throughout the infection thread. (E) Dual infection threads inside a root hair. The threads began as jumbled-type threads but later gave rise to green sectors at their tips (arrows show the sectors in each thread). (Reprinted from reference 60.)

View larger version (36K): [in this window] [in a new window]   FIG. 6. Model of infection thread growth. (A). A computer model was used to simulate sectored infection thread growth. For these cases, it was assumed that the thread started with six alternating bright and dark sectors of equal size. The doubling time for the growing bacteria (those 60 µm or less from the growing tip of the thread) was 4 h. Each bar is a representation of the length of the thread and its six sectors at each time point. Growth of the thread is exponential until it reaches 60 µm in length (dashed line) and becomes linear thereafter. Once the tipmost sector is more than 60 µm in length, it will be the only sector which continues to increase in size (see the last bar as an example). (B) Same as panel A, except the size of the growth zone is 20 µm. The thread grows at the same rate as for panel A while the thread is less than 20 µm long. After that, only cells in the tipmost 20 µm contribute to subsequent growth, and the overall rate of thread extension is lower than in the case where the growth zone is 60 µm long. (C). Same as panel A, except bacteria entering the thread are not assumed to have strictly alternated in terms of fluorescence. The entry pattern shown was dark-light-dark-dark-dark-light. This gives a total of four sectors, with the large dark sector in the middle of the thread having arisen from the loading of three dark cells in a row.

(ii) The model predicts that sectored infections should give rise to nodules that are populated with only one of the two infecting strains. This outcome occurs because the only subpopulation of bacterial cells which continues to advance down the infection thread indefinitely are the descendants of the bacteria which were at the very tip of the infection thread when it initiated.

(iii) The model predicts that growth of the bacterial population in the thread will be exponential while the infection thread is shorter than the growth zone. After that, growth of the bacterial population will be linear and the extension rate will be proportional to the size of the growth zone (Fig. 6B). This implies that the growth zone must relatively large in order to ensure quick transit of the infection thread through root tissue.

It is most parsimonious to assume that most infection threads, whether mixed or not, grow in the same manner as sectored threads, with bacteria near their tips actively growing, more distal bacteria not growing, and only descendants of the tipmost bacterium going on to populate the nodule. The random distribution of colored bacteria in some infection threads would appear to be in conflict with this idea and with the growth model outlined in Fig. 6. However, mixed infection threads in which the infecting strains are seemingly randomly distributed often become sectored after a time. This suggests that the growth patterns of these two types of threads are not too different. An addition to the model may explain the apparently random growth pattern seen in some threads (60).

It should be noted that the model outlined here holds when the increase in bacterial number inside infection threads is manifested as an increase only in the length, and not in the width, of the space occupied by the bacteria. This is the case for most infection threads that are actively growing down alfalfa root hairs. Infection threads developing in the infection zone of alfalfa nodules are much broader, and less uniform in width, than those seen in root hairs. The growth behavior of such bacterial populations is likely to be different than the growth behavior of the bacterial populations in the model above (Gage, personal observation) (Fig. 7).

View larger version (65K): [in this window] [in a new window]   FIG. 7. Architecture of the infection thread network inside a 10-day-old alfalfa nodule. An alfalfa nodule, induced by wild-type S. meliloti strain Rm1021, was fixed, embedded, and sliced into 1-µm-thick sections. Fifteen sections were dyed to reveal the bacteria and plant cell structures, photographed, and reassembled into a three-dimensional volume. (A) Projection of all 15 sections onto a single plane. The nuclei in each section are outlined in red, and the infection threads are filled in with green. The infection thread network appears to be polarized and growing toward the meristem, which is in the left side of the image. (B) Three-dimensional reconstruction of the nuclei and infection threads, showing a volume from the central part of the data set shown in panel A.

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The spatial localization of pericycle activation next to protoxylem poles may require an inducing factor, termed stele factor and thought to perhaps be uridine, coming from the protoxylem, or it may be caused by inhibition of the response of cells near the protophloem poles (100, 142, 151). mRNA encoding 1-aminocyclopropane-1-carboxylic acid (ACC) oxidase, which catalyzes the final step of ethylene synthesis, has been shown by in situ hybridization to be enriched in cells adjacent to the protophloem poles (74). Given that ethylene can inhibit nodulation at a variety of steps (see below), it may be that expression of ACC oxidase at the protophloem poles increases ethylene production there and prevents activation of pericycle cells and nodule formation at these sites. Of course, an activating factor such as uridine at protoxylem poles may act in concert with an inhibitor such as ethylene at protophloem poles to provide the spatial information that results in columns of activated cells aligned over the protoxylem poles (see Fig. 1B for an example).

Concomitant with the outward-propagating wave of cell activation, a column or two of cells in the outer cortex also begin to divide. The cytoplasm moves from the cell periphery to a central position, as it normally does during cell division, but the cells usually progress no further through the cell cycle (161, 169). The cytoplasms in these activated outer cortical cells align with each other, giving rise to columns of cytoplasmic bridges called preinfection threads (PITs) through which the inwardly growing infection thread propagates. T