Organelle Extensions in Plant Cells


  • Jaideep Mathur,

    Corresponding author
    1. Laboratory of Plant Development and Interactions, Department of Molecular and Cellular Biology, University of Guelph, 50 Stone Road, Guelph, Ontario N1G 2W1, Canada
    Search for more papers by this author
  • Alena Mammone,

    1. Laboratory of Plant Development and Interactions, Department of Molecular and Cellular Biology, University of Guelph, 50 Stone Road, Guelph, Ontario N1G 2W1, Canada
    Search for more papers by this author
  • Kiah A. Barton

    1. Laboratory of Plant Development and Interactions, Department of Molecular and Cellular Biology, University of Guelph, 50 Stone Road, Guelph, Ontario N1G 2W1, Canada
    Search for more papers by this author

Tel: +1 519 824 4120; Fax: +1 519 837 1802; E-mail:


Cell walls lock each cell in a specific position within the supra-organization of a plant. Despite its fixed location, each cell must be able to sense alterations in its immediate environment and respond rapidly to ensure the optimal functioning, continued growth and development, and eventual long-term survival of the plant. The ultra-structural detail that underlies our present understanding of the plant cell has largely been acquired from fixed and processed material that does not allow an appreciation of the dynamic nature of sub-cellular events in the cell. In recent years, fluorescent protein-aided imaging of living plant cells has added to our understanding of the dynamic nature of the plant cell. One of the major outcomes of live imaging of plant cells is the growing appreciation that organelle shapes are not fixed, and many organelles extend their surface transiently in rapid response to environmental stimuli. In many cases, the extensions appear as tubules extending from the main organelle. Specific terms such as stromules from plastids, matrixules from mitochondria, and peroxules from peroxisomes have been coined to describe the extensions. Here, we review our present understanding of organelle extensions and discuss how they may play potential roles in maintaining cellular homeostasis in plant cells.

  • image

[ Jaideep Mathur (Corresponding author)]


Plants and animals coexist in many different environments. Animals are able to physically locomote in response to changes in their immediate environment. However, plants adjust their pattern of growth and development according to the altered conditions of temperature, precipitation, wind, nutrients, and diverse biotic factors while remaining fixed in a particular location. Interestingly, a scaled-down version of the spatial relationship between a multi-cellular plant and the environment also exists for individual cells of a plant. Walls lock each plant cell in position within the supra-organization of the plant. From its singular location, each cell contributes toward the optimal functioning and long-term survival of the plant. Clearly, single cells can contribute meaningfully only when each of their sub-cellular components functions optimally. To achieve this optimal functioning, there must be mechanisms in place within the cell to minimize the negative effects of different kinds of environmental stresses to which the cell is exposed. Furthermore, as many stresses can rise rapidly to reach lethal levels, the adjustments within a plant cell must be made very quickly.

Although logic dictates that the plant cell must show dynamic internal behavior, the picture of the plant cell imprinted in our minds does not convey the impression of an active cell. This picture is based largely on transmission electron micrographs where the plant cell is seen to be comprised of discrete organelles and vesicles packed within thick walls. Although they preserve ultra-structural detail beautifully, electron micrographs completely erase the reality of dynamic organelle behavior that is characteristic of a living plant cell. The advent of time-lapse cine-microphotography, using normal transmitted light as well as differential interference contrast (DIC) microscopy techniques (Wildman et al. 1962; Green 1964; Allen and Allen 1983), changed this picture. Though conspicuously lacking the fine resolution of electron microscopy, these techniques provided an appreciation of subcellular motility. New insights on organelle motility suggested organelle interactions within the streaming cytoplasm (Wildman et al. 1962; Gunning 2005). A major wave of live imaging-based observations followed the cloning of the green fluorescent protein (GFP) (Prasher et al. 1992), and the targeting of it and other multi-color derivative fluorescent proteins to various subcellular components (reviewed by Tsien 1998; Mathur 2006; Shaw 2006; Sparkes and Brandizzi 2012). The strides in fluorescent protein-based labeling techniques have been matched by concomittant advances in laser-based optical imaging and data processing. Together, these techniques have opened a 4-dimensional (length -x, breadth -y, height -z, and time -t) window on the behavior of cytosolic fluids, membranes and membrane-bound organelles, and vesicles in living plant cells.

One of the major observations obtained through live imaging has been the visualization of dynamic extensions from different organelles in plant cells. As shown in this review, many of these extensions are formed in rapid response to environmental stimuli and help in maintaining homeostasis in plant cells.

The Plant's Perspective: Rapid Organelle Responses Differ from Long-term Responses of a Cell

Time-lapse imaging has made us aware of how quickly sub-cellular structures and fluids are moved around within living cells. Two categories of responses can be distinguished on the basis of the time required for their execution and the consequences of the response.

Organelle responses belonging to the first category are directly relevant to the survival of a single cell and its optimal functioning. These responses are rapid and occur within seconds or minutes of a cell perceiving a signal. More importantly, these responses are transient. They extend the interacting surface of an organelle greatly but do not bring about a change in the number of organelles of a specific kind within the cell. Depending upon the extent and intensity of the causal stress, the rapid subcellular response might occur in a small region of a cell, involve a single cell, or become spread out between groups of cells. Furthermore, the rapid subcellular responses might involve a single organelle such as a plastid, or several organelles such as plastids, mitochondria, and peroxisomes acting in concert. For many organelles, the rapid response to alterations in their microenvironment is manifested as a change in their motility, causing them to form large aggregates (Logan and Leaver 2000; Mathur et al. 2002). Alternatively, chloroplast response to quality and quantity of light (Oikawa et al. 2003), the nuclear response to pathogen invasion (Gross et al. 1993; Iwabuchi et al. 2007), and the ER and peroxisomal response to pathogens (Hardham et al. 2008) result in rapid displacement from one subcellular locale to another. However, the observed change in motility or its directionality for a particular organelle might indicate a more general involvement of several factors, including alterations in localized ionic gradients (Monshausen et al. 2009; Tanaka et al. 2010), cytoskeletal organization (Mathur et al. 2002; Akkerman et al. 2011; Hamada et al. 2012), and motor protein activity (Reisen and Hanson 2007; Verchot-Lubicz and Goldstein 2010). Rapid responses that can be directly associated with specific organelles involve the formation of one or more elastic tubular extensions from the main organelle body (Figures 1, 2). These responses are transient and are easily missed because the organelle reverts to its initial state very quickly after responding.

Figure 1.

Plastid extensions and their key features. 

(A) Chloroplasts (ch) and their stromules (st) differentially coloured using a photoconvertible stroma targeted tpFNR:mEosFP. All plastids including chloroplasts extend stromules sporadically. A large main plastid body (mb) containing thylakoid membranes arranged in grana can be distinguished from the thin tubular stromules extended by chloroplasts. Chlorophyll autofluorescence in this picture is depicted in blue while the green, red and intermediate hues are due to the mEos fluorescent protein. 
(B) Plastids (pt) that extend and retract stromules (st) frequently cluster around the nucleus (n). Nucleus is dyed red following staining with a vital SYTO-25 nucleic acid dye from Invitrogen-Molecular Probes. 
(C) As shown here, nuclei can also sporadically exhibit 5–15 μm long extensions (ne). Like stromules, nuclear tubules also exhibit extension and retraction with respect to the main nuclear body. 
(D) Extensive stromules observed in an arc5 mutant of Arabidopsis thaliana illustrate their alignment along stretches of RFP-highlighted ER tubules. Arrowheads point to stromule branch points that display a strong correlation with the branching of cortical ER tubules. 
(E) Transient bulges (arrowheads) are often observed along stromules. The inset ‘e’ shows a tubular leucoplast from an A. thaliana root cell that shows prominent domains reminiscent of bulges in ‘E’. Leucoplasts and other tubular plastids are pleomorphic, and such bulged domains are constantly being formed and reorganized. 
Ch, chloroplasts; er, endoplasmic reticulum tubules; mb, main body of a plastid; n, nucleus; ne, tubular extension from nucleus; pt, plastids; st, stromule; 
Size bars: E= 25 μm; all others 5 μm.

Figure 2.

Diagrammatic representation of mitochondrial and peroxisomal extensions and their relation to the division of the two organelles. 

(A) Peroxisomes extend and retract (grey arrows) thin peroxules (a) sporadically in response to increased subcellular ROS. Like stromules, the proxules exhibit bulged domains along their length. The bulged domains may be mistaken for a peroxisome ‘bud,’ but time-lapse observations usually show their retraction to the parent body. Peroxule formation does not result in an increase in peroxisome numbers. However, the grey rectangular background depicts a multi-gene regulated pathway that involves elongated peroxisomes and leads to an increase in peroxisomal numbers. A single peroxisome becomes elongated (b) through the coordinated activity of several genes. Elongated peroxisomes become beaded and undergo fission to produce more peroxisomes. Each new peroxisome is capable of forming peroxules. Peroxule extension and retraction is thus clearly different from the elongation and fission of peroxisomes. 
(B) Panel depicting mitochondria that might display thin tubular regions. Unlike plastids and peroxisomes, mitochondria undergo frequent fusion. Although not demonstrated in plants to date, matrixules might be involved in establishing the pre-fusion contacts between two mitochondria (a). Following fusion (b), a mitochondrion appears greatly elongated (c), and like all other elongated organelles exhibits narrow as well as dilated domains along its length. In the absence of knowledge about its internal structure, whether or not the narrow region (box with ‘?’) connecting two portions of an elongated mitochondrion represents a matrixule remains debatable. Division of mitochondria involves a multi-gene regulated machinery that produces many small mitochondria, each capable of fusing again.

In comparison to the rapid responses, the second response category requires a longer time for manifestation. Since the growth of a plant occurs through a combination of cell division and cell differentiation, these responses are tailored to create a particular pattern of growth and development. The simple division of a cell results in the creation of two daughter cells, each with its own complement of organelles. At the end of division, there is an overall increase in the number of organelles and other subcellular contents. The processes of organelle and cell division involve a large number of proteins and their precisely-coordinated interactions. In contrast, cellular differentiation does not increase the number of cells, but makes a cell different from its neighbors through various processes such as localized expansion, preferential and localized deposition of wall material, or removal of specific cellular components. The number of organelles within a cell may also change considerably during differentiation. Importantly, even under optimal conditions, both division and differentiation require hours or even days to reach completion.

Organelle extensions

Following advances in video enhanced cine-photography and confocal microscopy, the 4D-imaging of cells has clearly established that organelles extend their surface sporadically (Wildman et al. 1962; Menzel 1994; Köhler et al. 1997; Jedd and Chua 2002; Logan and Leaver 2000; Gunning 2005). However, live imaging of organelles is not without its own caveats. Tubular organelles in the cytoplasm can twist, roll and form loops within a matter of seconds. With each twist and turn, the profile of an organelle can change rapidly, and through bending and kinking a single long tubule can exhibit more than one profile simultaneously. Often, the changes are too quick to be recorded using the 3-dimensional x,y,z scan on a confocal laser scanning microscope, while x,y scans of the same plane over time lack the information from other planes. Nevertheless, it has been possible to convincingly document narrow diameter tubular extensions being extended and retracted by several organelles (Figures 1, 2).

For peroxisomes, mitochondria, and plastids, these intermittently-formed extensions have been called peroxules, matrixules, and stromules, respectively. In addition, assorted vesicles, vacuoles and the nucleus also produce tubular extensions sporadically (Verbelen and Tao 1998; Mathur et al. 2010; Sliwinska et al. 2012). Once extended by an organelle, the tubules might persist for a few hours if the conditions involved in their elicitation persist without becoming increasingly stressful (see subsequent discussion on stromules). Alternatively, the extensions might be retracted and re-extended in quick succession within a matter of seconds (for example, in the case of peroxules). The position from which an extension appears and the length and diameter of the extended tubules vary considerably over time. Many organelle extensions found in the cell cortex display branching. Nearly all tubules observed to date stretch and contract continuously and frequently display dilations along their length.

A major issue that can lead to confusing observations needs to be addressed before discussing organelle extensions further.

Elongated Organelles and Organelle Extensions are Different from Each Other

Diagrammatic representations of subcellular structures have created a typical picture of each organelle in our minds. Plastids are depicted as oval, mitochondria as bean-shaped, the nucleus usually spherical, and so on. However, time-lapse imaging shows that organelle shape is not fixed, and many organelles such as plastids and mitochondria exhibit a high degree of pleomorphy. Indeed, based on comparisons with the typical shape, a number of mutants showing aberrant organelle morphology have been isolated in Arabidopsis (Table 1). In addition, certain exogenously-applied treatments can also lead to changes in organelle morphology (Table 2). The terms peroxules, for peroxisomal extensions, and matrixules, for extensions from mitochondria, were introduced by Scott et al. (2007) on the basis of their strong morphological resemblance to stromules, the stroma-filled tubules extended by plastids (Köhler et al. 1997; Köhler and Hanson 2000). However, as discussed in subsequent sections dealing with stromules, peroxules and matrixules, a purely morphological resemblance is insufficient to differentiate between different elongated organelles and sporadically-formed organelle extensions.

Table 1.  Some proteins whose altered expression levels cause extension or elongation of organelles in Arabidopsis thaliana
Organelle geneAltered expression level due toPhenotypePostulated mechanismReferences
 AtFtsZ1-1Overexpression1-2 severely elongated chloroplasts per cellInhibition of chloroplast division Stokes 2000
 ARTEMISMutation/Anti-senseTriangular and elongated chloroplastsComponent for cell and chloroplast division Fulgosi et al. 2002
 ARC3, ARC6, ARC5MutationGiant chloroplasts that fail to divide efficiently; greatly increased stromulesIncomplete chloroplast division apparatus Pyke 1999; Holzinger et al. 2008
 AtCDT1RNAiEnlarged chloroplasts, cell division slowedInteracts with ARC6 to complete the chloroplast division apparatus Raynaud et al. 2005
 ATG5MutationIncreased stromule abundance and length during starvation conditionsAutophagosome pathway disrupted; causes stress / loss of membrane sequestration Ishida et al. 2008
 MSL2-1 MSL3-1Double mutantEnlarged plastids with no stromules. A few stromules after 15–30 minutes with 30% sorbitol. Many stromules after 60 minute of plant dehydration.Plastid membranes lack postulated mechanosensitive ion channels which contribute to maintaining plastid size and shape Veley et al. 2012
 PGM1MutationElongated plastids with underdeveloped internal membranesDefective in sugar – starch metabolism Casper et al. 1985; Schattat et al. 2012a.
 DRP3A and DRP3BDominant negative mutant proteinElongated mitochondriaEssential to the mitochondrial division apparatus Arimura et al. 2002
 BIGYIN 1-2MutationEnlarged mitochondriaComponent of the mitochondrial division apparatus, a FIS1 homolog Logan 2006; Zhang and Hu, 2008
 NMT1/ ELM1MutationElongated mitochondria; reticulumRequired for localization of DRP3A to the mitochondrial division site Logan et al. 2003; Logan 2006; Arimura et al. 2008
 DRP3A/APM1MutationElongated peroxisomes and mitochondriaRequired for peroxisome and mitochondrial division apparatus Mano et al. 2004; Fujimoto et al. 2009
 FIS1A FIS1BFIS1A mutation combined with FIS1B RNAiEnlarged peroxisomes and mitochondriaRequired for peroxisome and mitochondrial division apparatus Zhang et al. 2009; Zhang et al. 2008
 PEX11 c,dOverexpressionElongated peroxisomesPeroxisome membrane proteins that promote elongation, aggregation and division Lingard and Trelease 2006; Orth et al. 2007
 PEX3-1 PEX3-2iDouble knockdown RNAiElongated peroxisomesRequired to maintain the structure of the peroxisome. Nito et al. 2007
 DRP5B (ARC5)MutationClustered peroxisomes showing membrane constriction but impaired divisionInvolved in the peroxisome and chloroplast division apparatus Zhang et al. 2010
Table 2.  Some exogenous treatments and conditions that result in alterations in organelle behaviour including the formation of organelle extensions
Treatment or conditionPhenotypePostulated mechanismReferences
Related to tissue/organ specific development
Creation of cell suspensions“Millipede or octopus-like” extremely elongated plastids (Tobacco)Immature plastids with increased stromules; high sucrose and phytohormone media. Kohler and Hanson 2000.
Dark grown BY2 cellsElongated plastidsPlastids import sugars Schattat and Klösgen 2011
Creation of callusElongated plastids and increased stromule abundance. Branched networks of plastidsIncreased stromules in immature plastids, high sucrose and phytohormone media. Kohler and Hanson 2000
Normal mature mesophyll and trichomesDecreased stromule abundanceAdditional plastid contact with the cytosol is not required due to high chloroplast density. Plastids are fully developed. Waters et al. 2004
Normal Non-photosynthetic cells of epidermis, petals, rootsIncreased stromule abundanceIncreased stromules in immature plastids. Etioplasts and non-green plastids may respond with increased stromules for functions separate from photosynthesis. Shiina et al. 2000; Holzinger et al. 2007; Kohler et al. 1997, Kohler and Hanson 2000
Ripe fruitIncreased stromule abundance (Tomato)Low chloroplast density in non-green tissue triggers increase in plastid surface area for increased contact with cytosol. Waters et al. 2004
Normal cells in Alpine plantsThicker stromulesGenetic or adaptive plastid response to colder climate conditions. Lutz and Engel 2007
Related to disruption in organelle motility
Cytochalasin D, amiprophosmethylDecreased stromule abundance (Tobacco)Direct inhibition of actin and tubulin based motility, respectively. Kwok and Hanson 2003
2,3-butanedione monoximeLoss of stromule movement, decrease in stromule length and abundance (Tobacco)Directly inhibits actin-myosin motility at the myosin ATPase inhibitor. Natesan et al. 2009
Response to temperature
Temperature fluctuationUndetectable stromules at 10 °C, many stromules at 40 °CHigher temperatures are associated with more fluid membranes. Holzinger et al. 2007a; Buchner et al. 2007
Response to low oxygen concentration
2.4 kPa oxygenElongated mitochondriaHypoxia stresses mitochondria and may trigger their division. Ramonell et al. 2001
Anoxia for more than 4 hoursElongated and fused mitochondria (Tobacco cell culture)Mitochondrial fusion as a stress response in order to optimize recombination of mitochondrial DNA. Gestel and Verbelen 2002
Response to osmotic stress
Vacuum infiltration of 40 mM sucrose or glucose solution into leavesIncreases stromule abundance – sugar analogues or sugar metabolism mutants may produce a similar effectChloroplasts must import and store sugars as cellular sugar concentration increases. Schattat and Klösgen 2011
500 mM mannitol solutionIncreased stromules and some filamentous chloroplasts in leaves (Wheat)Osmotic stress triggers stromule formation. Gray et al. 2012
≥ 200 mM potassium chlorideOver 80% of chloroplasts emitting stromules (Tobacco)Salt stress triggers stromule formation. Gray et al. 2012
100 μM abscisic acidOver 80% of plastids emitting stromules (Tobacco)Mimics salt stress and dark growing conditions, dependent on cytosolic protein synthesis. Gray et al. 2012
6% polyethylene glycolOver 80% of plastids emitting stromules (Tobacco)Stimulated drought conditions at the plastid trigger stromules. Gray et al. 2012
Leaf dried at room temperature for 4 hoursIncreased stromule abundance (Tobacco)Desiccation triggers stromule formation. Gray et al. 2012
Response to alternative oxidase induction
Antimycin A (50 and 10 μM)Extreme, reversible plastid elongation in root cortexTreatment blocks the mitochondrial bc1 complex in the electron transport chain, triggering the alternative oxidase pathway, independent of protein synthesis. Respiration may be reduced causing increased cellular sugar concentration. Itoh et al. 2010; Itoh and Fujiwara 2010
RotenoneIncreased stromule abundanceInhibits complex I in the mitochondrial electron transport chain. Respiration may be reduced causing increased cellular sugar concentration. Itoh et al. 2010
Response to reactive oxygen species
0.08 to 0.8 M H2O2Significant increase in peroxules, cessation of motility after long treatmentsHydroxyl stress increases subcellular demand for peroxisomal catalase. Can induce peroxisomal fission. Sinclair et al. 2009
15-90 second UV-A treatmentElongated peroxisomesProduces subcellular H2O2, increases subcellular demand for peroxisomal catalase. Can induce peroxisomal fission. Sinclair et al. 2009
Cells undergoing arbuscule formation during mycorrhizae colonizationIncreased chloroplasts and stromule abundance with localization to the nucleus (Tobacco)Subcellular demand for sugars increases triggering plastid response. Cell invasion also triggers ROS accumulation, peroxisomes may respond. Fester and Hause 2005; Fester et al. 2001.
100 μM s-triazineTriggers elongated mitochondriaProduces superoxide during the day by blocking the chloroplast electron transport chain at photosystem II. Scott and Logan 2008; Bliek 2009
100 μM methyl vlologenTriggers elongated mitochondriaProduces superoxide radicals that can trigger apoptosis. Scott and Logan 2008; Bliek 2009
Pathogen attack mimic response
1-aminocyclopropane-1-carboxylic acid (ACC)Increased stromule abundanceMimics pathogen attack conditions when the cell produces ethylene (ACC is the first precursor in the committed ethylene synthesis pathway). Gray et al. 2012
1μM silver nitrateDecreased stromule abundanceInhibits ethylene activity. Gray et al. 2012

Elongated plastids should not be confused with stromules

The term “stromule” was coined to describe narrow, 0.3 to 0.8 μm diameter stroma-filled tubules ranging in length from a few micrometers to nearly 50–70 μm, that are extended by all kinds of plastids (Köhler et al. 1997; Köhler and Hanson 2000). In both electron micrographs (Lütz and Engel 2007; Holzinger et al. 2008; Sage and Sage 2009) and epi-fluorescent microscopy (Kohler and Hanson 2000; Gray et al. 2001), the tubular stromules can be clearly distinguished from the main spheroidal plastid body of chloroplasts, which contains well organized stacks of membrane thylakoids arranged in grana (Buchanan et al. 2000; Figure 1A). The tubular stromules are easily observed from plastids that are aggregated around nuclei (Figure 1B, C). However, the simple definition of a stromule as “a stroma-filled tubule” does not emphasize its extension from an independent plastid. With this definition, any stroma-filled region of a plastid with an elongated, tubular profile would qualify as a stromule. This does not pose a problem for chloroplast stromules since there is a clear and visible difference between the spherical main body containing grana and the narrow tubule which lacks organized internal membranes (Figure 1A). However, when one considers plastid types, such as leucoplasts in root cells, etioplasts in dark-grown cells and seedlings, confusion can arise. These plastids are tubular (Kirk and Tilney-Bassett 1978; Buchanan et al. 2000; Schattat et al. 2012), as they posses loosely-arranged, pro-lamellar bodies and thin, internal membrane tubules instead of the well organized thylakoid stacks that are typical of chloroplasts. If judged on a morphological basis, they resemble stromules. In addition, as observed for nearly all tubules including stromules, these elongated plastids display contraction and extension as well as occasional branching (Figure 1D; Schattat et al. 2012). Often, these plastids display no clear morphological equivalent of the main plastid body (Figure 1E, e) to serve as a reference point for demarcating an extended stromule. Furthermore, to make the observations even more confusing, the elongated plastids exhibit bulged domains sporadically along their length (Figure 1E; Pyke and Howells 2002). The presence of several of these bulged regions, which are sometimes large enough to suggest a plastid body, in an elongated tubule, give the appearance of several interconnected plastids. However, time-lapse imaging often shows the merging and reformation of such bulged domains (Schattat et al. 2011b; 2012).

One unfortunate and unintended consequence of the simple definition of a stromule (Köhler and Hanson 2000) is that tubular leucoplasts and etioplasts can sometimes have been mis-interpreted as stromules. Indeed, the seminal publication on the re-discovery of stromules entitled “Exchange of protein molecules through connections between higher plant plastids” (Köhler et al. 1997), presented the localized two-photon aided photo-bleaching of GFP in tobacco root plastids (Köhler et al. 1997). The tubular root leucoplasts with bulged domains shown in the figure were interpreted as two plastids interconnected by a stroma-filled tubule, ostensibly a stromule, and GFP was shown to flow within this compartment. This suggested that plastids interconnected by a tubule are able to exchange fluorescent proteins and, by extension, other macromolecules. Notably, later observations on stromules extended by independent plastids that remained in close proximity without exchanging photo-bleached GFP allowed Köhler and Hanson (2000) to conclude that exchange of plastid contents might not be a primary function of stromules. However, while minimizing the possibility of an interconnected plastid network for exchanging proteins, the authors maintained that the reason for not observing such protein exchange between chloroplasts in their experiments was probably because the stromules had not fused with one another (Köhler and Hanson 2000). Their observations left open the possibility that stromules could actually connect plastids. Despite the clear experiments of Hanson and Kohler (2000), a slight mis-representation of facts and speculations crept into reviews (Gray et al. 2001; Kwok and Hanson 2004a) and textbooks (Buchanan et al. 2000; Hanson and Köhler 2006; Evert 2006) that stromules might mediate inter-plastidic exchange of macromolecules. Unfortunately, early studies on stromules did not attempt to confirm whether the organelles being considered as “interconnected plastids” had ever been independent organelles at all before they apparently became connected through a tubule. It was assumed that the stroma-filled regions represented stromules, and therefore the areas that were connected must represent independent plastids.

Recently, Schattat et al. (2012a) reported the critical reappraisal of plastid interconnectivity and the possibility of fluorescent protein exchange between plastids. A green-to-red photo-convertible monomeric EosFP marker protein (Mathur et al. 2010) was used to differentially color individual plastids and their stromules (Figure 1A) before assessing their connectivity. Several experiments carried out on different kinds of plastids in wild type and arc5 and arc6 mutant plants of Arabidopsis expressing the plastid stroma targeted EosFP have strongly reinforced the independent nature of plastids (Schattat et al. 2012a). The observations indicate that contrary to reports based on apparently interconnected plastids (Köhler et al. 1997; Kwok and Hanson 2004a), large macromolecules such as fluorescent proteins may not be exchanged through the stromules sporadically extended by independent plastids. While using a different class of fluorescent proteins, Schattat et al. (2012) actually reaffirmed the conclusions reached by Köhler and Hanson (2000). The non-involvement of stromules in exchanging plastid DNA and plastid ribosomes has been observed independently by Newell et al. (2012). Furthermore, Schattat et al. (2012b) have demonstrated that when plastids such as etioplasts with differently-organized internal membranes undergo division, they stretch out considerably and appear as tubules containing two or more bulged regions (Figure 1E, e). During these stages, the narrow stroma-filled isthmus that connects the two halves of a dividing plastid (Waters and Pyke 2005) becomes readily visible. Despite giving the impression of connecting two plastids, the isthmus of an elongated dividing plastid should not be equated with stromules extended and retracted by independent plastids. Based on their observations, Schattat et al. (2012b) recommended that in order to avoid future confusion between elongated and dividing plastids, the use of the term “stromule” should be limited to extensions formed from independent plastids. Until it is unequivocally demonstrated that individual plastids can actually fuse with each other, a stroma-filled tubule stretching between regions of a single dividing or elongating plastid should be called an isthmus (Schattat et al. 2012a, 2012b). We suggest that this convention for stromules and isthmus should be maintained.

While stromules form the best-characterized organelle extensions and probably have important roles in maintaining cellular homeostasis (see later section on possible roles of organelle extensions), peroxules provide another example of rapid subcellular responses.

Peroxules and elongated peroxisomes

In an interphase plant cell, peroxisomes are accepted as spherical, approximately 0.8 to 1.5 μm diameter organelles that, amongst other functions, are intimately involved in the scavenging of hydrogen peroxide (H2O2) and other harmful reactive oxygen species (ROS; de Duve and Baudhuin 1966; Tabak et al. 2006). In plants, peroxisomes exhibit acto-myosin-based motility along the general cytoplasmic streaming that takes place within a cell (Jedd and Chua 2002; Mathur et al. 2002; Mano et al. 2002; Collings et al. 2002). Peroxisome numbers within a cell increase in response to a variety of stresses (Palma et al. 1991; Schrader et al. 1999; Frohnmeyer and Staiger 2003; Sinclair et al. 2009). As part of their proliferation response, single peroxisomes undergo elongation into 3 to 7-μm long tubules (Figure 2; Tabak et al. 2006; Titorenko and Mullen 2006). At least five PEROXIN11 proteins (Pex 11 a–e) are implicated in the elongation of peroxisomes (Lingard and Trelease 2006; Orth et al. 2007; Pan and Hu 2011). Subsequently, the vermiform, elongated peroxisome undergoes fission through the activity of at least three dynamin-related proteins DRP3A, DRP3B and DRP5B, and the FISSION1A and FIS1B proteins (Zhang and Hu 2010). Mutations in the DRP3A/B and FIS1A/B genes in Arabidopsis exhibit abnormally elongated peroxisomes that are unable to efficiently undergo fission (Zhang and Hu 2010; Hu et al. 2012).

In addition to the elongation and pre-fission beaded state, single peroxisomes also display thin projections that extend and retract from the spherical peroxisomal body (Figure 2A; Sinclair et al. 2009; Mathur 2009). Tubular extensions were described earlier for several of the torus marker lines generated through random GFP:cDNA fusion, and following immuno-localization with anti-catalase antibodies were considered to be peroxisomes (Cutler et al. 2000). Observations of transgenic plants expressing GFP with a carboxy terminal peroxisome targeting sequence (PTS1) also showed thin tubules being extended by peroxisomes. The presence of a fluorescent knob at their end led to the suggestion that new peroxisomes might bud off from elongated tubules produced by mature peroxisomes (Jedd and Chua 2002).

All these observations of peroxisomal extensions can now be grouped under the name “peroxules” (Scott et al. 2007). Peroxules up to 8 μm long have been observed from peroxisomes in cells exposed to transient increases in light intensity and other conditions such as exposure to H2O2 that lead to a rise in the subcellular level of hydroxyl stress (Sinclair et al. 2009; Mathur 2009). In a manner reminiscent of stromules, peroxules also extend, retract, and display bulged domains along their length. However, peroxule formation is limited to a short time window since prolonged exposure of a plant cell to intense light or to H2O2 inhibits the elastic behaviour of peroxules and instead results in the formation of single, 3–5 μm long vermiform peroxisomes (Sinclair et al. 2009). Upon returning to more moderate conditions, the vermiform peroxisomes break up into smaller peroxisomes, each of which is capable of extending peroxules again. The transient formation of peroxules thus clearly precedes the elongation of a single peroxisome. The change from a peroxule-extending peroxisome (Figure 2A-a) to an elongated peroxisome (Figure 2A-b) provides insight into the manner in which the plant cell responds and handles stress thresholds.

Apparently, subcellular ROS below a certain threshold lead to peroxule formation, whereas higher levels of the same stress results in an increased number of peroxisomes in the cell. Though peroxules can display strong fluorescence in any region along their length to suggest the presence of a small peroxisome (Jedd and Chua 2002), the actual breaking of a “bud” from peroxules to increase the peroxisomal population has not been observed.

Further elucidation of a “stress threshold sensing mechanism” to inform us of the difference between peroxule extension versus complete elongation of a peroxisome is provided by the aberrant peroxisome morphology (apm1) mutant in the dynamin-related protein 3A (DRP3A) gene of Arabidopsis (Mano et al. 2004). Peroxisomes in drp3A/apm1 mutants are enlarged, and their numbers per cell are reduced. During seedling development, these peroxisomes become abnormally elongated due to an apparent inability to carry out the fission step efficiently (Mano et al. 2004). However, even though the elongated peroxisomes in the apm1 mutants look and behave very much like stromules, based on a morphological criterion they could be called peroxules (Scott et al. 2007). Therefore, an alternative interpretation of the phenotype is that it might depict a prolongation of the peroxule phase.

Elongated peroxisomes, but not peroxules, are also obtained upon the altered expression of a number of other peroxisomal membrane proteins (Table 1), such as PEX10 (Scott et al. 2007), PEX11 isoforms (Lingard and Trelease 2006; Orth et al. 2007), and PEX16 (Karnik and Trelease 2005, 2007). A closer appraisal of the peroxisome phenotype in relevant mutants and in over-expression and RNAi lines (Nito et al. 2007; Hu et al. 2012) would be greatly instructive in our understanding of the relative roles that stress thresholds might play in the formation and maintenance of organelle extensions.

Matrixules and elongated mitochondria

Mitochondria present yet another aspect of organelle extension. Generally depicted as 0.5 to 5 μm long, oval- to sausage-shaped, double-membrane bound compartments, mitochondria are amongst the most pleomorphic organelles in plant cells (Diers 1966). The changes in their morphology are rapid, and are accompanied by rounds of fission and fusion (Logan and Leaver 2000; Arimura et al. 2008; Schwarzländer et al. 2012). Their fusion can result in up to 80 μm-long chains that form a tangled mitochondria reticulum (Segui-Simarro and Staehelin 2009). Alternatively, anoxia induces mitochondrial expansion and results in giant flattened disc-shaped mitochondria (Van Gestel and Verbelen 2002). A number of mitochondrial mutants that show aberrant aggregation and odd, greatly elongated morphology have been isolated in Arabidopsis (Logan et al. 2003; Arimura et al. 2008; Table 1).

Like peroxules, the term matrixules was coined for extensions from mitochondria that look similar to plastid stromules (Scott et al. 2007). However, it is unknown whether or not matixules only contain the “matrix,” like stromules which are known to contain only the stroma. Like stromules from independent plastids, and in accordance with the clarifications provided earlier in this review, the term “matrixules” should be strictly applied only to sporadic extensions from single mitochondria. Although elongated mitochondria occur frequently, we have been unable to find descriptions of such mitochondrial extensions in plants. However, a singular report by Bowes and Gupta (2008) describes thin tubular extensions from mitochondria in animal cells from the B-SC-1 African green monkey kidney. These extensions were observed approximately 25 min after exposing the cells to N-ethylmaleimide (NEM), and appeared just before mitochondrial fusions took place. Since this report did not explore whether the extensions were filled only by the matrix or also contained infolded cristae, it would be inappropriate to label them as matrixules. Because mitochondrial fusion takes place routinely in plant cells too (Figure 2B; Arimura et al. 2004) in response to different conditions and chemicals (Table 2), it would be interesting to see whether similar thin tubular extensions are also observed just before mitochondrial fusion.

The observation that mitochondrial fusion and fission occur routinely creates an interesting problem from the perspective of imaging matrixules in living cells. During fusion, two mitochondria (Figure 2B-a) merge together to form a longer mitochondrion (Figure 2B-b, c;Schattat et al. 2012a). The long organelle obtained through fusion is already tubular, and often stretches further to display thin regions within the tubule (Diers 1966). Mitochondrial chains made of many fused mitochondria also display multiple similarly thin, tubular regions as they stretch and contract while moving as part of the general cytoplasmic stream. Whether or not thin portions of a tubular mitochondrion should be considered matrixules remains debatable. For peroxules and stromules, an additional criterion has been applied under which they are considered to be extensions, since their formation does not lead to an increase in their population. The rapid fusion and fission of mitochondria does not allow this criterion to be applied in assessing matrixules, since in a living, rapidly-responding plant cell it is never clear how many single mitochondria actually exist.

Elongated mitochondria are also observed in addition to elongated peroxisomes in the apm1/drp3a mutants (Arimura et al. 2002; Logan et al. 2004). Plastids in these mutants also extend stromules, and thus provide a unique system to understand the similarities and differences between tubular organelles and organelle extensions.

Other organelles also display extensions

Live imaging-based observations on vacuolar compartments show the occurrence of extensive tubules or “ripples” on the vacuolar surface (Verbelen and Tao 1998; Kutsuna et al. 2003; Reisen et al. 2005; Wiltshire and Collings 2009). Electron micrographs and schematics based on these ripples show long tubules from pro-vacuoles, and suggest that the progressive fusion and expansion of tubules is responsible for the formation of large vacuoles (Marty 1978). Indeed, recent observations on transgenic plants over-expressing a photo-convertible mEosFP fused to the PI3P-specific 2xFYVE domain show vacuoles that have a 10–20 μm diameter that extend and retract tubules of diameters varying from 250 nm to 1 μm (Mathur et al. 2010). Currently, it is unknown if tubules from vacuoles occur under certain conditions such as irradiation by intense light, as suggested by the observations of Wiltshire and Collings (2009), or if they are artifacts associated with the over-expression of PI3P-binding domains like the FYVE domain (Mathur et al. 2010).

Recent observations on the movement of nuclei during tip growth of root hairs and collet hair cells in Arabidopsis show nuclear extensions that appear as tubules squeezing past large vacuoles (Sliwinska et al. 2012). Like peroxules, these nuclear extensions are extended and retracted, and often change the direction of extension. However, following their retraction, the nuclear shape reverts to its characteristic spherical-to-oval shape. Similar transient nuclear extensions have been observed within plastid clusters around the nucleus (Figure 1C). At this stage, it is unclear which components of the nucleus are actually present in these extended organelles. Furthermore, because the observations were largely based on confocal microscopy, it is not clear whether they represent real tubules or simply appear tubular due to the profile of a flattened nucleus squeezing past a large turgid vacuole.

Conditions eliciting organelle extensions suggest their different roles in cellular homeostasis

In agreement with the transient nature of organelle extensions, it seems reasonable to suggest that the phenomenon is regulated by recurrent but short-lived changes within the cell's micro-environment. Indeed, experiments carried out to elicit peroxules used exogenous H2O2 application or varied the intensity and time at which cells were exposed to light (Sinclair et al. 2009). Under these conditions, peroxules were formed and retracted within a matter of minutes. Since one of the major roles ascribed to peroxisomes is the scavenging of subcellular ROS, including H2O2 (de Duve and Baudhuin 1966), peroxules might have a role in transiently increasing the interactive membrane surface between peroxisomes and the cytosol. This might create an efficient scavenging system that quickly removes the ROS from the cell before they reach toxic levels.

In comparison to peroxules, stromules can remain extended for long periods of time, although they might exhibit variation in their length and diameter during extension. Many conditions, including altered subcellular redox status (Itoh et al. 2010), symbiotic interactions (Fester et al. 2001; Hans et al. 2004; Lohse et al. 2005), elevated temperatures (Holzinger et al. 2007a), viral infection (Caplan et al. 2008), and changes in plastid size and population in a cell (Pyke and Howells 2002; Waters et al. 2004) have been implicated in the extension of stromules. While each of these situations creates substantial changes in the cellular environment, two sets of observations published recently suggest a strong correlation between stromule extension and the levels of sugars within a cell. Exogenous feeding of sucrose to epidermal cells was shown to result in a discernable increase in stromule extension frequency from epidermal plastids (Schattat and Klösgen 2011c). In a follow-up study, the frequency of stromule formation was found to fluctuate between day and night (Schattat et al. 2012b). In general, only a few leaf epidermal plastids in Arabidopsis plants extended stromules at the end of a dark period. However, exposure to light for as little as 3 h caused the stromule extension frequency in a cell to rise many fold, and then to stay constant at a high level for as long as the light persisted. A return to darkness resulted once more in retraction of the stromules. Extended dark periods maintained a low frequency of stromule extension, while extended light consistently kept the stromule frequencies in a high range (Schattat et al. 2012b). These new observations on the diurnal rhythm of stromule formation are exciting, as they suggest that stromules might be integral components of the machinery involved in maintaining sugar metabolism within a cell. In this context, it is interesting to note that in wheat amyloplasts, protrusions contain small starch granules that suggest a role for stromules in sugar-starch related metabolic phenomena (Langeveld et al. 2000).

Stromules have also been considered to be interaction platforms for mitochondria and peroxisomes (Kwok and Hanson 2004b; Gunning 2005; Holzinger et al. 2007a, 2007b). While most respiratory inhibitors do not induce changes to plastid morphology, antimycin-A triggers extreme, reversible filamentation in root cortex plastids of Arabidopsis (Itoh et al. 2010). Antimycin-A activity is independent of cytoskeletal regulation and protein synthesis, but may be associated with the alternative oxidase electron transport pathway in mitochondria combined with an interorganellar signal (Itoh et al. 2010). Several other roles for stromules have been suggested in many reviews (Gray et al. 2001; Natesan et al. 2005; Lopez-Juez and Pyke 2005; Hanson and Sattarzadah 2011), and their validation requires further experimentation.

The published literature provides a number of abiotic and biotic conditions under which mitochondrial morphology changes (Table 2). However, in many cases, the time involved in response elicitation has not been given due consideration. Additionally, as stated in the section on matrixules, their actual relation to the main body of the mitochondrion is not always clear. Therefore, more detailed investigation on matrixules is required before any conjecture on their possible roles can be made. A similar situation exists for vacuoles and nuclei, where the current amount of information is still very limited. In general, extensions are expected to increase the interactivity of an organelle with other organelles and the cytosol. Investigations involving the endoplasmic reticulum (ER) and the actin cytoskeleton are particularly interesting in this context.

Contiguous regions of the endoplasmic reticulum appear to be involved in shaping the morphology and behavior of extensions from other organelles

Numerous transmission electron micrographs depict close associations between the ER and other organelles such as plastids, mitochondria, peroxisomes, the nucleus, and vacuoles (Diers 1966; Crotty and Ledbetter 1973; Gibbs 1979; Ehara et al. 1985; McLean et al. 1988; Bourett et al. 1999; Holzinger et al. 2007a, 2007b; Lutz and Engel 2007; Velikanov et al. 2011a, 2011b). While the outer nuclear envelope is known to be continuous with the peri-nuclear ER, mitochondria-associated ER membrane (MAM) sites form contact points between mitochondria and the endoplasmic reticulum (Hayashi et al. 2009; Giorgi et al. 2009; Friedman et al. 2011). Similar membrane contact sites (MCSs) called plastid associated membranes (PLAM) have been described for plastids (Andersson et al. 2007). Physical association between the ER and the PLAM has been demonstrated on isolated pea (Pisum sativum) chloroplasts using optical imaging and laser tweezers, whereby pulling forces of up to 400 pN could not detach ER fragments attached to the plastid envelope, suggesting strong protein-protein interactions at these points (Andersson et al. 2007). The case for connectivity between the ER and the peroxisomes has been debated for more than half a century, and is still open to further investigation (Hu et al. 2012). Nevertheless, both stromules (Schattat et al. 2011a, 2011b; Figure 1D) and peroxules (Sinclair et al. 2009; Mathur 2009) have been observed aligning with contiguous ER tubules. In the case of cortically-located stromules extended by epidermal plastids, branching occurs at angles that match the angles created by polygons of tubular ER (Schattat et al. 2011; Figure 1D). In addition, simultaneous observation of GFP-highlighted stromules and RFP-labeled ER show that stromules extend and retract within ER-lined channels in a manner dictated by ER organization (Schattat et al. 2011a, 2011b). In a similar manner, peroxule extension occurs in patterns dictated by contiguous ER tubules (Sinclair et al. 2009).

Whether or not the dynamic relationship observed between stromules and the ER and between peroxules and the ER actually results from MCSs between the organelles remains to be seen. Since ER-PLAMs might be directly involved in lipid trafficking, it is interesting that the transient expression of a Brassica napuschloroplast lipase protein (BnCLIP1) fused to a GFP localizes as small punctae on the chloroplast outer membrane. When BnCLIP1:GFP was expressed simultaneously with an ER targeted yellow fluorescent protein (YFP), the punctae appear to mark the sites where ER tubules contact the plastid (Tan et al. 2011). Notably, in transient over-expression experiments, a few of the BnCLIP:GFP punctae appear to localize away from the chloroplasts, and might represent MCSs with other organelles or the plasma membrane (Tan et al. 2011). Alternatively, these punctae could be localized on extended stromules. Discrete foci have also been observed on chloroplasts in transgenic plants expressing mechano-sensitive-ion channel-like (MSL) proteins MSL2 and MSL3 fused to different fluorescent proteins (Haswell and Meyerowitz 2006). Notably, an Arabidopsis double mutant lacking both MSL 2 and 3 exhibits large round epidermal plastids that lack dynamic stromules. This phenotype is present under normal growth conditions, and does not require exposure to extracellular osmotic stress. However, the epidermal plastids in msl2-1 msl3-1 leaves exhibit rapid and reversible changes in their volume and shape in response to exogenously-applied hypertonic or hypotonic challenges (Veley et al. 2012). The proposed link between cellular osmotics and the extension of stromules opens an exciting avenue for exploration.

Organelle extensions and their association with the actin cytoskeleton

In plants, nearly all organelle movement appears to rely upon the actin-myosin system, and ER organization and behaviour have been directly associated with actin dynamics and F-actin organization (Quader et al. 1987; Kachar and Reese 1988; Lichtscheidl and Url 1990; Sparkes et al. 2009; Yokota et al. 2011). It can therefore be argued that the alignment of organelle extensions with ER tubules is because both the ER and the organelle membrane associate with the same F-actin strand or bundle. As suggested by Schattat et al. (2011a, 2011b) this may indeed be the case, since different motor molecules, each carrying their own unique cargo, can employ a single cytoskeletal track for movement.

Chloroplasts are surrounded by baskets of actin filaments (Kandasamy and Meagher 1999), and a close association between stromules and actin microfilaments has been suggested (Kwok and Hanson 2004; Gunning 2005). When tobacco hypocotyls were treated with actin inhibitors such as cytochalasin D or latrunculin B, there was a change in both plastid morphology and plastid movement (Kwok and Hanson 2003). Furthermore, long stromules were lost upon drug treatment, and arc-like structures that appeared to be retracted stromules that had lost tension were observed (Kwok and Hanson 2003). Gray et al. (2001) also reported the cessation of stromule dynamics in tobacco (Nicotiana tabacum) and onion (Allium cepa) cells upon treatment with latrunculin B, while a similar observation was made for stromules in leaf mesophyll cells of Oxyria digyna (Holzinger et al. 2007b).

The link between acto-myosin and stromules has been further strengthened through observations on a myosin XI motor (Natesan et al. 2009). While YFP fusion with the entire tail region of myosin XI-F localizes primarily to the cytoplasm, its truncated tail domain fusion with YFP was found to localize to plastids and stromules upon transient expression in N.benthamiana (Sattarzadeh et al. 2009). Notably, mutant lines with insertions in all 13 of the known myosin XI genes in Arabidopsis do not exhibit any defects in chloroplast movement (Peremyslov et al. 2008).

For peroxules, further investigation is required, since extensions were sometimes observed in areas that did not explicitly exhibit an F-actin strand. Furthermore, F-actin organization did not change when approximately 1.2 μm diameter spherical peroxisomes were made to stretch out into 3–6 μm long tubules through 60 s of violet-blue irradiation (Sinclair et al. 2009). These observations might be imaging artifacts, since it is extremely difficult to image fine F-actin. A myosin XI isoform has been shown to localize to peroxisomes (Hashimoto et al. 2005), and fluorescent protein fusion of the tail region of several myosin-XI genes has been observed to localize to small organelles ranging in size from 0.5–3 μm (Reisen and Hanson 2007). Whether or not a specific myosin localizes to peroxisomes and dictates peroxule activity is yet to be determined.

Organelle Extensions Play an Important Role Within the Overall Survival Strategy of a Plant

The enlargement of an organelle to a particular size, the triggering of different protein cascades leading to the recruitment of specific proteins to mediate its division, and the actual process of division all require time. In contrast, the formation of organelle extensions is rapid and transient. Apparently, such extensions can minimize subcellular stress levels efficiently without having to add more organelles to the cell. While serving an important house-keeping function, the formation of organelle extensions also indicates that the living plant cell recognizes stress thresholds and responds appropriately. When the stress levels in a cell exceed a particular threshold, the molecular machinery for organelle multiplication is triggered. A direct demonstration of this is suggested through observations on peroxisomes where peroxules form an intermediate state that can lead to peroxisome elongation and fission (Figure 2A-b), or revert back to a single, parental peroxisome (Figure 2A-a). In turn, the increased number of organelles within a cell may determine the preparedness of a cell for further differentiation, or lead to its division. Thus, within the context of the bigger strategy for plant survival, the rapid responses of single organelles can be thought of as small checkmarks whose frequent occurrence paves the way for more complex but relatively long-term responses within the plant cell. Discovering the common mechanisms underlying rapid organelle extension promises an exciting foray into the fundaments of the eukaryotic cell.

(Co-Editor: Jianping Hu)


We would like to thank Cristina Ruberti for her critical comments. Research funding by the Natural Sciences and Engineering Research Council of Canada (NSERC), the Canada Foundation for Innovation (CFI), and the Ministry of Research and Innovation, Ontario, is gratefully acknowledged.