Plant organelles predominantly rely on the actin cytoskeleton and the myosin motors for long-distance trafficking, while using microtubules and the kinesin motors mostly for short-range movement. The distribution and motility of organelles in the plant cell are fundamentally important to robust plant growth and defense. Chloroplasts, mitochondria, and peroxisomes are essential organelles in plants that function independently and coordinately during energy metabolism and other key metabolic processes. In response to developmental and environmental stimuli, these energy organelles modulate their metabolism, morphology, abundance, distribution and motility in the cell to meet the need of the plant. Consistent with their metabolic links in processes like photorespiration and fatty acid mobilization is the frequently observed inter-organellar physical interaction, sometimes through organelle membranous protrusions. The development of various organelle-specific fluorescent protein tags has allowed the simultaneous visualization of organelle movement in living plant cells by confocal microscopy. These energy organelles display an array of morphology and movement patterns and redistribute within the cell in response to changes such as varying light conditions, temperature fluctuations, ROS-inducible treatments, and during pollen tube development and immune response, independently or in association with one another. Although there are more reports on the mechanism of chloroplast movement than that of peroxisomes and mitochondria, our knowledge of how and why these three energy organelles move and distribute in the plant cell is still scarce at the functional and mechanistic level. It is critical to identify factors that control organelle motility coupled with plant growth, development, and stress response.

Plant cells are often envisioned as the classic textbook cartoon: static organelles spaced indiscriminately in the cytoplasm within the confines of rigid cell walls. However, organelles are highly dynamic, as their quantities fluctuate via biogenesis, fission, fusion, and degradation and their morphologies shift among a variety of sizes and shapes. Organelles traverse the cell by cytoplasmic streaming and organelle-specific, targeted motility along the cytoskeleton, driven by molecular motors that travel along the tracks of the cytoskeletal filaments.

Eukaryotic cells employ the cytoskeletal motors of kinesins, dyneins, and myosins that hydrolyze ATP to propel organelle transport directionally along polarized microtubule and actin tracks, whose orientation dictates the direction of travel. Active motor-mediated movement can trigger other modes of transportation, including cytoplasmic streaming and ‘hitchhiking', in which organelles are transported indirectly by associating with cargoes already being carried by motors [1].

In animal cells, the microtubule-based cytoskeleton, along with kinesin and dynein motors, are predominantly responsible for vesicle and organelle trafficking [2–4]. In contrast, actin filaments comprise the major intracellular highway system in plants for long-range organelle trafficking, driven primarily by the class XI myosin proteins [5–9]. Plant microtubules and their associated motors, kinesins, mainly direct short-range organelle movement, pauses, and orientation in coordination with the actin cytoskeleton [10–14]. Additionally, the endoplasmic reticulum (ER) is intricately involved in motor-assisted organelle motility. For example, the ER is associated with the dynamic cytoskeletal network through tethering and anchoring mechanisms, most especially at ER-plasma membrane contact sites [6,15]. Furthermore, proximity and interactions among organelles correlate with fluctuations in ER morphology, which suggests that the ER mediates the positioning and movements of organelles, including chloroplasts, mitochondria and peroxisomes [16,17].

A number of studies testing organelle motility in Arabidopsis myosin mutants and myosin localization in plant cells indicate functional redundancy in the Myosin XI protein family and targeting of the same myosin motor to multiple types of organelles [5,18–22]. The apparent promiscuity in the interactions between myosin motors and organelles beckons for regulatory mechanisms of organelle movement that is possibly governed by organelle-specific receptors/adaptors, which recruit designated Myosin XI motors [23].

These molecular motors and their associated cytoskeleton facilitate the movement and dynamics of the nucleus and endomembrane system, including ER, Golgi, and vesicles, which have been expertly reported in several recent papers [24–27]. In this review, we focus on the motility of plant energy organelles — chloroplasts, mitochondria, and peroxisomes — which work independently as well as in concert during energy metabolism. These organelles modulate not only their metabolism and morphology but also their distribution and motility in accordance with developmental stages and environmental conditions. Here, we introduce the collaborative nature of these three energy organelles in photosynthetic organisms and methods used to study their motilities and discuss the factors that trigger and modulate their movement and distribution. We apologize to researchers whose publications cannot be cited due to space limitation.

Chloroplasts, enclosed by double envelopes and containing their own genome, house the photosynthetic machinery that performs light capture and carbon assimilation, along with many other vital cellular functions beyond direct energy production, such as the biosynthesis of lipids, phytohormones and other key molecules [28–31]. Mitochondria are double-membraned, DNA-containing organelles whose primary role is cellular respiration while also performing plant-specific functions such as malate oxidation and photorespiration [32]. Peroxisomes are single-membraned and DNA-less organelles involved in fatty acid degradation and detoxification of reactive oxygen species, along with plant-specific functions such as phytohormone production and photorespiration [33–35].

Several of the above-mentioned processes require the collaboration of chloroplasts, mitochondria, and/or peroxisomes [36–38]. Namely, the photorespiratory pathway requires these three organelles to recover mis-assimilated carbon produced due to the oxygenase activity of the photosynthetic enzyme Rubisco, protecting the plant cell from toxic byproducts [39,40]. Besides being essential to the survival of organisms performing oxygenic photosynthesis in ambient air, photorespiration also has a demonstrated role in immunity [41]. In addition, during the metabolism of fatty acids, triacylglycerol stored in oil bodies can be degraded in peroxisomes via β-oxidation, after which the glyoxylate cycle, also housed in peroxisomes, produces metabolites that are exported to mitochondria for the tricarboxylic acid (TCA) cycle to ultimately release the stored energy [33,42]. Moreover, lipid transfer can occur directly between chloroplasts and mitochondria during phosphate starvation [43]. As a final example, jasmonate (JA) biosynthesis requires chloroplasts to produce the precursor 12-oxo-phytodienoic acid (OPDA), which is then imported into the peroxisome to be converted to JA via β-oxidation [33,44,45].

Consistent with their metabolic connections, these organelles are often observed to be in close physical proximity. Mitochondrial associations with chloroplasts increase under high light in the diatom Phaedactylum, as well as during mixotrophy (compared with phototrophy) in the alga Nannochloropsis [46]. In Arabidopsis, the proximity area between these organelles expands in light compared with dark [47]. Chloroplast-peroxisome-mitochondrion complexes are formed more frequently in light than in dark, and light-induced triple organelle interaction is suggested to be regulated by photosynthesis and facilitated by ER-chloroplast nexuses [47–49]. ER reorganization is also proposed to contribute to the formation of tubular protrusions from chloroplasts, peroxisomes, and mitochondria, termed stromules, peroxules, and matrixules, respectively [50]. During high light irradiation, small mitochondria cluster around peroxules [49]. While proximity among all three energy organelles increases in mesophyll in light [47,48] and stromule formation is more frequent in light-treated epidermal pavement cells (Figure 1A), similar numbers of mitochondria and peroxisomes interact with both the chloroplast body and stromules in these pavement cells [51]. These organelle protrusions very likely provide a platform for inter-organellar interaction and metabolite exchange among the organelles.

Examples of energy organelle dynamics in photosynthetic organisms during environmental and developmental changes.

Figure 1.
Examples of energy organelle dynamics in photosynthetic organisms during environmental and developmental changes.

Changes in the distribution and movement of chloroplasts/plastids (green), mitochondria (yellow), and peroxisomes (pink) are shown. (A) Chloroplasts and mitochondria exhibit photorelocation and cold positioning in Arabidopsis and tobacco pavement cells (PC), mesophyll cells (MC), and guard cells (GC). Peroxisome proliferation and motility increase in light. Light-induced morphological changes include mitochondrial aggregation, peroxisome elongation, and chloroplast stromule formation. (B) Seasonal rearrangement of chloroplasts occurs in conifer species. (C) Chloroplasts redistribute during pathogen infection and immune response in Arabidopsis and tobacco leaves. (D) The three energy organelles are differentially distributed and vary their motility during pollen tube development in flowering plants. (E) Dynamics of the three energy organelles shift in response to reactive oxygen species (ROS) in Arabidopsis leaves. Mitochondria slow and aggregate while peroxisomes speed up and form peroxules. (F) Mitochondria slow down, redistribute under fluctuating carbon dioxide (CO2) levels in Chlamydomonas. (G) Peroxisomes are more concentrated in the meristematic zone in root tips and their morphology is altered in response to cadmium (Cd2+) toxins in roots. Arrows in (A), (D) and (E) represent general motility patterns. MeJA, methyl jasmonate; NO, nitric oxide. The nucleus (blue) is only depicted when its interaction with energy organelles is involved. Created with BioRender.com.

Figure 1.
Examples of energy organelle dynamics in photosynthetic organisms during environmental and developmental changes.

Changes in the distribution and movement of chloroplasts/plastids (green), mitochondria (yellow), and peroxisomes (pink) are shown. (A) Chloroplasts and mitochondria exhibit photorelocation and cold positioning in Arabidopsis and tobacco pavement cells (PC), mesophyll cells (MC), and guard cells (GC). Peroxisome proliferation and motility increase in light. Light-induced morphological changes include mitochondrial aggregation, peroxisome elongation, and chloroplast stromule formation. (B) Seasonal rearrangement of chloroplasts occurs in conifer species. (C) Chloroplasts redistribute during pathogen infection and immune response in Arabidopsis and tobacco leaves. (D) The three energy organelles are differentially distributed and vary their motility during pollen tube development in flowering plants. (E) Dynamics of the three energy organelles shift in response to reactive oxygen species (ROS) in Arabidopsis leaves. Mitochondria slow and aggregate while peroxisomes speed up and form peroxules. (F) Mitochondria slow down, redistribute under fluctuating carbon dioxide (CO2) levels in Chlamydomonas. (G) Peroxisomes are more concentrated in the meristematic zone in root tips and their morphology is altered in response to cadmium (Cd2+) toxins in roots. Arrows in (A), (D) and (E) represent general motility patterns. MeJA, methyl jasmonate; NO, nitric oxide. The nucleus (blue) is only depicted when its interaction with energy organelles is involved. Created with BioRender.com.

Close modal

Chloroplasts, due to their superiorly large size and chlorophyll autofluorescence, can be readily visualized through light and fluorescence microscopy [52].

In early studies, detections of peroxisomes and mitochondria were largely dependent on organelle-specific enzymes or distinct properties of their membranes. For example, using fluorescence microscopy, plant peroxisomes were detected by immunofluorescence using antibodies against catalase [53] while mitochondria were probed by dyes like rhodamine 123 that binds to P-glycoproteins [54,55]. Due to disadvantages associated with rhodamine 123 like challenges of sample washes and photostability, it has largely been replaced by synthetic hydrophobic dyes commonly known as MitoTrackers, which bind to mitochondrial membranes and can be conjugated with different fluorophores [56] to track mitochondria in living plant cells [57].

Lately, fluorescent protein (FP) tags introduced by recombinant DNA techniques have been widely adopted to monitor the dynamic behavior of peroxisomes and mitochondria, when either stable transformation or transient expression is applicable in the host cells. Besides the founding green fluorescent protein (GFP), numerous FPs may be chosen nowadays for their features of color, brightness, quantum yield, maturation property, pKa, and lifetime, which are frequently updated by the research community (https://www.fpbase.org/). When covalently attached to proteins that are unique to the respective organelle at either the organelle membranes or lumens after translation, these FPs faithfully report the intracellular positions of the organelles by fluorescence microscopy. Moreover, organelle and cytoskeletal FP marker collections have expanded beyond model plants and are now optimized for uses in crop plants like maize [58], rice [59,60], and barley [61]. Several excellent articles have summarized typical examples [62–64].

To label peroxisomes, a 3-amino acid canonical peroxisomal targeting signal 1 (PTS1) sequence, such as Ser-Lys-Leu (SKL) or Ser-Arg-Leu (SRL), is usually translationally conjugated to the C-terminus of an FP [63,65]. Peroxisomes can also be illuminated using the fusion of the N-terminus of a peroxisome matrix protein containing the PTS2 nonapeptide, such as that from the 3-ketoacyl-CoA thiolase or the glyoxysomal malate dehydrogenase, and the FP [66]. When an FP-PTS1 or PTS2-FP fusion protein is expressed under constitutively active promoters like the viral 35S promoter, it is in high abundance so that peroxisomes are brightly labeled for convenient observation by confocal microscopy.

Similarly, mitochondria can be labeled by attaching their targeting sequence at the N-terminus of FPs. A widely used targeting sequence in plants is a 29-amino acid peptide derived from the yeast cytochrome c oxidase IV (COX4) [63]. Mitochondrial-specific proteins like the γ-subunit of F1-ATPase serve the purpose equally well in plant cells [64]. Although they require transcription and translation in host cells, such FP-based probes are often favored over lipophilic dyes because of their organelle specificity.

The motility as well as polarized distribution of plant organelles including peroxisomes and mitochondria is dependent primarily on actin microfilaments and employs the myosin motors [18,53]. Live-cell imaging of actin filaments has evolved from the application of the fluorescent dye-conjugated fungal toxin of phalloidin to FP-tagged polypeptides of various actin-binding proteins [67,68]. Earlier adoption of the actin-binding protein talin often resulted in the formation of thick actin bundles [68]. As a replacement, ABD2, an actin-binding domain of the Arabidopsis fimbrin 1 protein, or the 17-amino acid Lifeact peptide derived from the yeast actin-binding protein Abp140 [69], is often used these days, both of which can significantly reduce the bundling and aggregation of actin microfilaments.

The ever-growing diversity of FPs and highly sophisticated microscopes with exceptional resolution enable synchronous visualization of multi-organellar and cytoskeletal topology in living cells. For example, peroxisomes, mitochondria, and actin labeled with multicolored FP-fusion biomarkers as described above, along with autofluorescent chloroplasts, can be monitored simultaneously in living tobacco epidermal cells (Figure 2). This concurrent visualization enables direct comparisons of organelles in the context of one another. Moreover, intricate visualization techniques have vast potential to further illuminate organelle dynamics and motility by rapid capture of organelles in action (Supplementary Movie 1).

Simultaneous visualization of plant energy organelles and actin in tobacco cells.

Figure 2.
Simultaneous visualization of plant energy organelles and actin in tobacco cells.

Chloroplasts (chlorophyll autofluorescence, green), mitochondria (COXIV29aa-eYFP, yellow), peroxisomes (mScarlet-I-SRL, red), and actin (Lifeact-eGFP, purple) in Nicotiana tabacum were imaged using a 3i spinning disk confocal microscope with excitation wavelengths 640 nm, 515 nm, 561 nm, and 488 nm, respectively. The emission capture range for each fluorophore is as follows: 672–712 nm for chlorophyll, 528.5–555.5 nm for eYFP, 580.5–653.5 nm for mScarlet-I, and 510–540 nm for eGFP. Scale bar represents 10 µm.

Figure 2.
Simultaneous visualization of plant energy organelles and actin in tobacco cells.

Chloroplasts (chlorophyll autofluorescence, green), mitochondria (COXIV29aa-eYFP, yellow), peroxisomes (mScarlet-I-SRL, red), and actin (Lifeact-eGFP, purple) in Nicotiana tabacum were imaged using a 3i spinning disk confocal microscope with excitation wavelengths 640 nm, 515 nm, 561 nm, and 488 nm, respectively. The emission capture range for each fluorophore is as follows: 672–712 nm for chlorophyll, 528.5–555.5 nm for eYFP, 580.5–653.5 nm for mScarlet-I, and 510–540 nm for eGFP. Scale bar represents 10 µm.

Close modal

FP-tagged probes are also used to detect the dynamic microtubule network in plant cells. The microtubule-binding domain (MBD) derived from the mammalian microtubule-associated protein 4 (MAP4) decorates cortical microtubules in fava bean epidermal cells upon transient expression [70]. This GFP-MBD renders superior brightness, possibly resulted from induced microtubule bundling. Alternatively, an MBD derived from the atypical casein kinase CKL6 has also been used to mark microtubules in living plant cells [71]. To minimize the potential impact of ectopically expressing an MBD in plant cells, one wishes to fuse FP with tubulins so that microtubules are labeled when the FP-tubulin fusion is incorporated into polymerized microtubules, which often requires stable transformation as demonstrated first by a GFP-α-tubulin in Arabidopsis [72]. The β-tubulin isoform TUB6 was later chosen because of its minimal toxicity after ectopic expression [73]. Stable expression of TUB6 under its native promoter in Arabidopsis results in neglectable disturbance to the dynamic remodeling of microtubule arrays and plant growth [74].

Progressively advanced, specific, and tunable visualization techniques equip researchers to uncover the nuances of energy organelle dynamics and expand our current knowledge of chloroplastic/plastidial, mitochondrial, and peroxisomal distribution and motility.

As light-harvesting apparatus, chloroplasts optimize photon capture by shifting their positions in the cell, a process termed photorelocation [75–78] (Figure 1A). In the dark, chloroplasts are distributed along the bottom of mesophyll cells and the top of pavement cells [51,79]. However, in shaded or low-light environments, chloroplasts accumulate across the peri-clinal cortex to increase light absorbance and energy capture, and pavement cell chloroplasts relocate to the bottom of the cell nearer to mesophyll chloroplasts. Conversely, when light intensity exceeds the plant's photosynthetic capacity, chloroplasts reduce photodamage by employing an avoidance mechanism in which they stack along the anti-clinal walls of the cell [51,80] (Figure 1A). During blue-light avoidance, nuclei move concordantly with chloroplasts by hitchhiking as passengers of chloroplasts to protect DNA from UV damage [81,82] (Figure 1A). While photorelocation is predominantly described in mesophyll cells where photosynthesis is most active, evidence suggests that chloroplasts are repositioned in an actin-dependent manner in guard cells in the epidermis as well. Shifts in light modulate stomatal aperture, which is also regulated by phytohormones produced in the chloroplast. Chloroplasts in tobacco guard cells redistribute from the inner peri-clinal walls to the dorsal walls at the center of guard cells during white light-induced stomatal opening (Figure 1A), a process proposed to be mediated by stochastic dynamics of actin filaments [83,84].

Because light availability fluctuates, chloroplasts must readily initiate photorelocation routinely. Blue-light receptors (phototropins) trigger the reorganization of specialized chloroplast actin filaments (cp-actin) via actin-bundling proteins, while Chloroplast Unusual Positioning Protein 1 (CHUP1) mediates interactions among chloroplasts, cp-actin, and the plasma membrane [77,79,85–87]. Although some data suggests the involvement of myosin proteins in chloroplast photorelocation [88], the process predominantly depends on actin reorganization itself in which cp-actin aggregates at the leading edge of chloroplasts to direct movement towards anti- or peri-clinal cell walls [83,87,89].

Critical to the cold acclimation of photosynthesis, the chloroplast avoidance response can be triggered by weaker light intensities at low temperatures in Arabidopsis (Figure 1A), suggesting the modulation of photoprotective mechanisms upon the onset of dark when temperatures drop [90,91]. Since cold positioning was first described in ferns [92], researchers have further elaborated on the links between photorelocation and cold acclimation in bryophytes [90,91] and flowering plants [90]. Phototropins act as thermosensors via blue-light regulated autophosphorylation [95] and subcellular re-localization [96–98], modulating the actin-dependent repositioning of chloroplasts as unfused aggregates in response to cold [99–101]. Seasonal rearrangement of chloroplasts was also observed in the mesophyll of two conifer species, in which chloroplasts move from the periphery along the cell wall in the summer to a more internal location in the cell in the winter (Figure 1B), a process that involves the vacuole and cytoplasmic strands [102].

Chloroplasts cluster around the nucleus during innate immune response. Oppositional movement of nuclei and chloroplasts is proposed to contribute to the formation of stromules enriched in proximity to the nucleus (Figure 1C), possibly to facilitate the exchange of defense signals [103–107]. Independent of nuclear movement, chloroplasts accumulate at the pathogen interface (Figure 1C) during Phytophthora infestans infection in Nicotiana benthamiana [108]. They even employ some components of the photorelocation mechanism for motility in Arabidopsis epidermal cells during fungal infections [109,110]. Immunity-related repositioning observed in tobacco is directed by the coordinated efforts of actin, microtubules, and stromules [103], all of which have dynamic roles in chloroplast motility and environmental stress response [111].

During pollen tube elongation, plastids exhibit differential motility patterns that are loosely correlated with a variety of plastid morphologies, where spherical plastids correspond with less streaming and elongated plastids and stromules with higher rates of streaming. Specifically, plastids move long-distances from pole-to-pole at high velocities (∼1.5 µm/s), with slower motions (∼0.4 µm/s) and arrests predominating in the middle region of the tube, and short vibrations or fluctuations at the tip end of the shank with a velocity of ∼0.5 µm/s [112,113] (Figure 1D).

Early characterizations of mitochondrial dynamics describe the organelle behavior and morphology as highly heterogeneous [114], which is further emphasized by the mitochondrial diversity recently observed across microalgae [46], indicating its multipotent functions in photosynthetic organisms. Mitochondria traverse plant cells along actin filaments in a variety of patterns, including short oscillations, bidirectional long-range movement, and turnaround motions facilitated by circular actin bundles [115,116]. Their morphology and motility are influenced by development, cell-type, proximity to other organelles, and environmental conditions, as described below.

The fundamental regulation of mitochondrial movement is likely combinatorial and collaborative, involving actin and myosins for various speeds and patterns of motion as well as microtubules and likely kinesins for positioning [116,117]. This translocation and positioning infrastructure enables the mitochondria to meet the energy demands of the cell. For example, lily mitochondria are concentrated at the sub-apex of the pollen tube (Figure 1D), where energy is required for elongation [14,118,119]. The pollen tube exhibits a characteristic reverse fountain-like pattern of cytoplasmic streaming in which organelles, including mitochondria, move through the shank towards the apex (∼2.67 µm/s) but then reverse course (Figure 1D) as the actin structure shifts from long filaments to shorter parallel bundles referred to as the actin fringe [118,120,121]. This fast forward movement followed by slowing down in the sub-apical region is attributed to actin/myosins and microtubules/kinesins, respectively [122]. In fact, specific motors Kinesin-like Protein 1 [123] and Myosin XI-C2 are associated with mitochondria, and Myosin XI-C2 contributes to mitochondrial as well as peroxisomal and Golgi movement in Arabidopsis pollen tubes [124].

ROS accumulation acts as a trigger for mitochondrial redistribution during heat stress-, UV- and methyl jasmonate (MeJA)-induced programmed cell death, in which mitochondria aggregate in clusters, shift from elongated morphology to more swollen and spherical shapes, and decrease their overall motility [125–127] (Figure 1E). These morphological changes are believed to be pivotal early steps in the progression of programmed cell death.

The trend towards more spherical, shorter, or simplified shapes for mitochondria is also evident in the transition from dark to light in Arabidopsis mesophyll cells [47] (Figure 1A). Light conditions also influence mitochondrial distribution and mobility. In Arabidopsis mesophyll cells, weak and strong blue light illuminations cause differential localization of mitochondria compared with mesophyll cells in the dark, mirroring chloroplast accumulation and avoidance [128] (Figure 1A). Mitochondrial long-range motility is dependent on actin, whereas they become more static in the proximity of chloroplasts, adopting slower, actin-independent wiggling-type motions [129]. In the green alga Chlamydomonas, mitochondria are closely associated with the plasma membrane in ambient CO2 and more uniformly distributed throughout the cytoplasm under high CO2 [130] (Figure 1F). These mitochondrial movements may facilitate necessary associations among chloroplasts, mitochondria, as well as peroxisomes during processes requiring their collaboration, such as photorespiration that is modulated by light and CO2 levels [131].

Despite having a simple structure, peroxisomes are highly dynamic in size, number, morphology, and biochemistry [33]. For instance, light increases peroxisome abundance in dark-grown Arabidopsis seedlings (Figure 1A) through a phytochrome A-mediated signaling pathway involving the transcription factors Homolog (HYH) and (Forkheaded-Associated Domain 3) FHA3 and the peroxisome elongation factor Peroxin 11b (PEX11b) [132–134]. In addition, peroxisomes in Arabidopsis mesophyll are significantly larger in light compared with the dark, and as their volume increases, they become less spherical [47] (Figure 1A). Moreover, chloroplast-associated peroxisomes are more elliptical (Figure 1A), increasing surface area available for inter-organellar interaction. This notion is further supported by stronger adhesion between these organelles in red/blue light, although these changes are not regulated by the red and blue light receptors phytochromes and phototropins [48]. In Arabidopsis root tips, peroxisomes are more intensely concentrated in the elongation and meristematic zones compared with the maturation zone, which is correlated with the finding that nitric oxide (NO), which exists in the peroxisome, distributes mostly in the primary and lateral root apices [135] (Figure 1G). Interestingly, cadmium (Cd2+)-imposed ROS increases peroxisome proliferation in Arabidopsis leaves and induces peroxule formation [136,137] (Figure 1E), while Cd2+ decreases the number of peroxisomes in primary roots and increases peroxisome size (Figure 1G), suggesting increased organelle fusion [135].

Peroxisomes display various patterns of movement: vibrations or oscillations in place, short-range travel, and traversing longer distances [7]. While myosin drives fast and long-range movement [18,138,139], oscillatory patterns of peroxisome motion correlate with ER dynamics in Arabidopsis seedling epidermal cells, suggesting a possible concomitant regulation of ER-peroxisome motility [140]. Moreover, individual peroxisomes switch directions, speeds, and motions all within the same cell type and timeframe in Arabidopsis epidermal cells [7]. As such, peroxisomes are not simply passive passengers but rather are subject to yet undiscovered biological triggers that govern their movements.

Developmental stage and tissue type is suggested to have an impact on peroxisome motility. Brownian-type vibrations have been ascribed to peroxisomes in leaves, with slower velocities in younger leaves, whereas more rapid peroxisomal movement is described in Arabidopsis roots, trichomes, mature leaves, and pollen tubes [19,121,138,139,141]. However, these velocities are highly variable: <0.2 µm/s in seedling epidermis [141], ∼2.23 µm/s in pollen tubes [121], up to 7 µm/s in seedling mesophyll [49], indicating that careful consideration regarding age, tissue, and cell type must be taken when evaluating peroxisome movements. Moreover, environmental factors likely influence peroxisomal motion. For example, significantly more peroxisomes are mobile in light compared with dark in Arabidopsis mesophyll cells (Figure 1A). Organelle mobility is maintained by actin filaments as peroxisome-chloroplast interactions increase, while peroxisome-chloroplast tethering is an actin-independent process involving peroxule formation [48,142]. Motility quickens during Cd2+-induced oxidative stress, which is hypothesized to be regulated by increased peroxisomal levels of ROS and may involve calcium signaling [141] (Figure 1E). While overall motility increases 1–3 days following Cd2+ treatment, peroxisomes were previously reported to pause during peroxule formation within 15 min of exposure to Cd2+, followed by an increase in proliferation [136,141]. These data suggest peroxisomes may dwell when undergoing morphological changes or intra/inter-organelle contact, and the number and size of peroxisomes may affect their motility.

  • Chloroplasts, mitochondria, and peroxisomes are essential organelles functioning independently as well as coordinately during plant energy metabolism and other key processes. The motility and distribution of these energy organelles are fundamentally important for plant physiology and defense.

  • Besides morphing between shapes, dividing and fusing, these energy organelles move throughout the cell and interact with one another and with other cell compartments to accomplish their roles efficiently. Most of what we know till now about their motility and distribution in photosynthetic organisms addresses general patterns and velocity changes in response to developmental and environmental cues, whereas the underlying mechanisms are largely unknown.

  • Many questions remain to be addressed in future research. For example, how are the molecular motors recruited to the organelles selectively? Which signaling pathways trigger the various modes of organelle positioning, movement, and physical interaction? What functional role does organelle motility play in plant physiology and health? How do these organelles coordinate their physical interaction and movement along the cytoskeletal tracks while performing collaborative metabolic functions?

The authors declare that there are no competing interests associated with the manuscript.

This work was supported by funding from The National Science Foundation to J.H. (MCB 2148206) and B.L. (MCB 2148207).

A.M.K., B.L. and J.H. co-wrote the manuscript. A.M.K. generated the figures.

CHUP1

Chloroplast Unusual Positioning Protein 1

COX4

cytochrome c oxidase IV

ER

endoplasmic reticulum

FP

fluorescent protein

GFP

green fluorescent protein

JA

jasmonate

MBD

microtubule-binding domain

MeJA

methyl jasmonate

NO

nitric oxide

PTS1

peroxisomal targeting signal 1

ROS

reactive oxygen species

SKL

Ser-Lys-Leu

SRL

Ser-Arg-Leu

1
Christensen
,
J.R.
and
Reck-Peterson
,
S.L.
(
2022
)
Hitchhiking across kingdoms: cotransport of cargos in fungal, animal, and plant cells
.
Annu. Rev. Cell Dev. Biol.
38
,
155
178
2
Olenick
,
M.A.
and
Holzbaur
,
E.L.F.
(
2019
)
Dynein activators and adaptors at a glance
.
J. Cell Sci.
132
,
jcs227132
3
Logan
,
C.M.
and
Menko
,
A.S.
(
2019
)
Microtubules: evolving roles and critical cellular interactions
.
Exp. Biol. Med. (Maywood)
244
,
1240
1254
4
Jongsma
,
M.L.M.
,
Bakker
,
N.
and
Neefjes
,
J.
(
2023
)
Choreographing the motor-driven endosomal dance
.
J. Cell Sci.
136
,
jcs259689
5
Sparkes
,
I.A.
,
Teanby
,
N.A.
and
Hawes
,
C.
(
2008
)
Truncated myosin XI tail fusions inhibit peroxisome, Golgi, and mitochondrial movement in tobacco leaf epidermal cells: a genetic tool for the next generation
.
J. Exp. Bot.
59
,
2499
2512
6
Perico
,
C.
and
Sparkes
,
I.
(
2018
)
Plant organelle dynamics: cytoskeletal control and membrane contact sites
.
New Phytol.
220
,
381
394
7
Mathur
,
J.
,
Mathur
,
N.
and
Hulskamp
,
M.
(
2002
)
Simultaneous visualization of peroxisomes and cytoskeletal elements reveals actin and not microtubule-based peroxisome motility in plants
.
Plant Physiol.
128
,
1031
1045
8
Ueda
,
H.
,
Tamura
,
K.
and
Hara-Nishimura
,
I.
(
2015
)
Functions of plant-specific myosin XI: from intracellular motility to plant postures
.
Curr. Opin. Plant Biol.
28
,
30
38
9
Nebenfuhr
,
A.
and
Dixit
,
R.
(
2018
)
Kinesins and Myosins: molecular motors that coordinate cellular functions in plants
.
Annu. Rev. Plant Biol.
69
,
329
361
10
Liang
,
Y.J.
and
Yang
,
W.X.
(
2019
)
Kinesins in MAPK cascade: how kinesin motors are involved in the MAPK pathway?
Gene
684
,
1
9
11
Ali
,
I.
and
Yang
,
W.C.
(
2020
)
The functions of kinesin and kinesin-related proteins in eukaryotes
.
Cell Adh. Migr.
14
,
139
152
12
Ali
,
I.
and
Yang
,
W.C.
(
2020
)
Why are ATP-driven microtubule minus-end directed motors critical to plants? An overview of plant multifunctional kinesins
.
Funct. Plant Biol.
47
,
524
536
13
Cai
,
G.
(
2022
)
The legacy of kinesins in the pollen tube 30 years later
.
Cytoskeleton (Hoboken)
79
,
8
19
14
Cai
,
G.
,
Parrotta
,
L.
and
Cresti
,
M.
(
2015
)
Organelle trafficking, the cytoskeleton, and pollen tube growth
.
J. Integr. Plant Biol.
57
,
63
78
15
Zang
,
J.
,
Kriechbaumer
,
V.
and
Wang
,
P.
(
2021
)
Plant cytoskeletons and the endoplasmic reticulum network organization
.
J. Plant Physiol.
264
,
153473
16
Mathur
,
J.
(
2020
)
Review: morphology, behaviour and interactions of organelles
.
Plant Sci.
301
,
110662
17
Mathur
,
J.
,
Kroeker
,
O.F.
,
Lobbezoo
,
M.
and
Mathur
,
N.
(
2022
)
The ER is a common mediator for the behavior and interactions of other organelles
.
Fron. Plant Sci.
13
,
846970
18
Avisar
,
D.
,
Prokhnevsky
,
A.I.
,
Makarova
,
K.S.
,
Koonin
,
E.V.
and
Dolja
,
V.V.
(
2008
)
Myosin XI-K Is required for rapid trafficking of Golgi stacks, peroxisomes, and mitochondria in leaf cells of Nicotiana benthamiana
.
Plant Physiol.
146
,
1098
1108
19
Prokhnevsky
,
A.I.
,
Peremyslov
,
V.V.
and
Dolja
,
V.V.
(
2008
)
Overlapping functions of the four class XI myosins in Arabidopsis growth, root hair elongation, and organelle motility
.
Proc. Natl Acad. Sci. U.S.A.
105
,
19744
9
20
Sattarzadeh
,
A.
,
Schmelzer
,
E.
and
Hanson
,
M.R.
(
2011
)
Analysis of organelle targeting by DIL domains of the Arabidopsis Myosin XI family
.
Front. Plant Sci.
2
,
72
21
Sattarzadeh
,
A.
,
Schmelzer
,
E.
and
Hanson
,
M.R.
(
2013
)
Arabidopsis myosin XI sub-domains homologous to the yeast myo2p organelle inheritance sub-domain target subcellular structures in plant cells
.
Front. Plant Sci.
4
,
407
22
Li
,
J.F.
and
Nebenfuhr
,
A.
(
2008
)
The tail that wags the dog: the globular tail domain defines the function of myosin V/XI
.
Traffic
9
,
290
298
23
Perico
,
C.
,
Gao
,
H.
,
Heesom
,
K.J.
,
Botchway
,
S.W.
and
Sparkes
,
I.A.
(
2021
)
Arabidopsis thaliana myosin XIK is recruited to the Golgi through interaction with a myoB receptor
.
Commun. Biol.
4
,
1182
24
Wada
,
M.
(
2018
)
Nuclear movement and positioning in plant cells
.
Semin. Cell Dev. Biol.
82
,
17
24
25
Kriechbaumer
,
V.
and
Brandizzi
,
F.
(
2020
)
The plant endoplasmic reticulum: an organized chaos of tubules and sheets with multiple functions
.
J. Microsc.
280
,
122
133
26
Renna
,
L.
and
Brandizzi
,
F.
(
2020
)
The mysterious life of the plant trans-Golgi network: advances and tools to understand it better
.
J. Microsc.
278
,
154
163
27
Zhang
,
R.
,
Xu
,
Y.
,
Yi
,
R.
,
Shen
,
J.
and
Huang
,
S.
(
2023
)
Actin cytoskeleton in the control of vesicle transport, cytoplasmic organization and pollen tube tip growth
.
Plant Physiol.
193
,
9
25
28
Sabater
,
B.
(
2018
)
Evolution and function of the chloroplast. Current investigations and perspectives
.
Int. J. Mol. Sci.
19
,
3095
29
Song
,
Y.
,
Feng
,
L.
,
Alyafei
,
M.A.M.
,
Jaleel
,
A.
and
Ren
,
M.
(
2021
)
Function of chloroplasts in plant stress responses
.
Int. J. Mol. Sci.
22
,
3464
30
Foyer
,
C.H.
and
Hanke
,
G.
(
2022
)
ROS production and signalling in chloroplasts: cornerstones and evolving concepts
.
Plant J.
111
,
642
661
31
Li
,
M.
and
Kim
,
C.
(
2022
)
Chloroplast ROS and stress signaling
.
Plant Commun.
3
,
100264
32
Moller
,
I.M.
,
Rasmusson
,
A.G.
and
Van Aken
,
O.
(
2021
)
Plant mitochondria - past, present and future
.
Plant J.
108
,
912
959
33
Pan
,
R.
,
Liu
,
J.
,
Wang
,
S.
and
Hu
,
J.
(
2020
)
Peroxisomes: versatile organelles with diverse roles in plants
.
New Phytol.
225
,
1410
1427
34
Hu
,
J.
,
Baker
,
A.
,
Bartel
,
B.
,
Linka
,
N.
,
Mullen
,
R.T.
,
Reumann
,
S.
et al (
2012
)
Plant peroxisomes: biogenesis and function
.
Plant Cell
24
,
2279
2303
35
Kaur
,
N.
,
Reumann
,
S.
and
Hu
,
J.
(
2009
)
Peroxisome biogenesis and function
.
Arabidopsis Book
7
,
e0123
36
He
,
C.
,
Berkowitz
,
O.
,
Hu
,
S.
,
Zhao
,
Y.
,
Qian
,
K.
,
Shou
,
H.
et al (
2023
)
Co-regulation of mitochondrial and chloroplast function: molecular components and mechanisms
.
Plant Commun.
4
,
100496
37
Gomez-Casati
,
D.F.
,
Barchiesi
,
J.
and
Busi
,
M.V.
(
2022
)
Mitochondria and chloroplasts function in microalgae energy production
.
PeerJ
10
,
e14576
38
Van Aken
,
O.
and
Van Breusegem
,
F.
(
2015
)
Licensed to kill: mitochondria, chloroplasts, and cell death
.
Trends Plant Sci.
20
,
754
766
39
Timm
,
S.
,
Florian
,
A.
,
Fernie
,
A.R.
and
Bauwe
,
H.
(
2016
)
The regulatory interplay between photorespiration and photosynthesis
.
J. Exp. Bot.
67
,
2923
2929
40
Sunil
,
B.
,
Talla
,
S.K.
,
Aswani
,
V.
and
Raghavendra
,
A.S.
(
2013
)
Optimization of photosynthesis by multiple metabolic pathways involving interorganelle interactions: resource sharing and ROS maintenance as the bases
.
Photosynth. Res.
117
,
61
71
41
Jiang
,
X.
,
Walker
,
B.J.
,
He
,
S.Y.
and
Hu
,
J.
(
2023
)
The role of photorespiration in plant immunity
.
Front. Plant Sci.
14
,
1125945
42
Li
,
N.
,
Xu
,
C.
,
Li-Beisson
,
Y.
and
Philippar
,
K.
(
2016
)
Fatty acid and lipid transport in plant cells
.
Trends Plant Sci.
21
,
145
158
43
Michaud
,
M.
,
Gros
,
V.
,
Tardif
,
M.
,
Brugiere
,
S.
,
Ferro
,
M.
,
Prinz
,
W.A.
et al (
2016
)
Atmic60 is involved in plant mitochondria lipid trafficking and is part of a large complex
.
Curr. Biol.
26
,
627
639
44
Pan
,
R.
,
Reumann
,
S.
,
Lisik
,
P.
,
Tietz
,
S.
,
Olsen
,
L.J.
and
Hu
,
J.
(
2018
)
Proteome analysis of peroxisomes from dark-treated senescent Arabidopsis leaves
.
J. Integr. Plant Biol.
60
,
1028
1050
45
Wasternack
,
C.
and
Strnad
,
M.
(
2018
)
Jasmonates: news on occurrence, biosynthesis, metabolism and action of an ancient group of signaling compounds
.
Int. J. Mol. Sci.
19
,
2539
46
Uwizeye
,
C.
,
Decelle
,
J.
,
Jouneau
,
P.H.
,
Flori
,
S.
,
Gallet
,
B.
,
Keck
,
J.B.
et al (
2021
)
Morphological bases of phytoplankton energy management and physiological responses unveiled by 3D subcellular imaging
.
Nat. Commun.
12
,
1049
47
Midorikawa
,
K.
,
Tateishi
,
A.
,
Toyooka
,
K.
,
Sato
,
M.
,
Imai
,
T.
,
Kodama
,
Y.
et al (
2022
)
Three-dimensional nanoscale analysis of light-dependent organelle changes in Arabidopsis mesophyll cells
.
PNAS Nexus
1
,
pgac225
48
Oikawa
,
K.
,
Matsunaga
,
S.
,
Mano
,
S.
,
Kondo
,
M.
,
Yamada
,
K.
,
Hayashi
,
M.
et al (
2015
)
Physical interaction between peroxisomes and chloroplasts elucidated by in situ laser analysis
.
Nat. Plants
1
,
15035
49
Jaipargas
,
E.A.
,
Mathur
,
N.
,
Bou Daher
,
F.
,
Wasteneys
,
G.O.
and
Mathur
,
J.
(
2016
)
High light intensity leads to increased peroxule-mitochondria interactions in plants
.
Front. Cell Dev. Biol.
4
,
6
50
Mathur
,
J.
(
2021
)
Organelle extensions in plant cells
.
Plant Physiol.
185
,
593
607
51
Barton
,
K.A.
,
Wozny
,
M.R.
,
Mathur
,
N.
,
Jaipargas
,
E.A.
and
Mathur
,
J.
(
2018
)
Chloroplast behaviour and interactions with other organelles in Arabidopsis thaliana pavement cells
.
J. Cell Sci.
131
,
jcs202275
52
Wada
,
M.
(
2013
)
Chloroplast movement
.
Plant Sci.
210
,
177
182
53
Collings
,
D.A.
,
Harper
,
J.D.I.
and
Vaughn
,
K.C.
(
2003
)
The association of peroxisomes with the developing cell plate in dividing onion root cells depends on actin microfilaments and myosin
.
Planta
218
,
204
216
54
Wu
,
F.S.
(
1987
)
Localization of mitochondria in plant-cells by vital staining with rhodamine-123
.
Planta
171
,
346
357
55
Gambier
,
R.M.
and
Mulcahy
,
D.L.
(
1994
)
Confocal laser-scanning microscopy of mitochondria within microspore tetrads of plants using rhodamine-123 as a fluorescent vital stain
.
Biotech. Histochem.
69
,
311
316
56
Poot
,
M.
,
Zhang
,
Y.Z.
,
Kramer
,
J.A.
,
Wells
,
K.S.
,
Jones
,
L.
,
Hanzel
,
D.K.
et al (
1996
)
Analysis of mitochondrial morphology and function with novel fixable fluorescent stains
.
J. Histochem. Cytochem.
44
,
1363
1372
57
Zheng
,
M.Z.
,
Beck
,
M.
,
Muller
,
J.
,
Chen
,
T.
,
Wang
,
X.H.
,
Wang
,
F.
et al (
2009
)
Actin turnover is required for myosin-dependent mitochondrial movements in Arabidopsis root hairs
.
PLoS ONE
4
,
e5961
58
Wu
,
Q.
,
Luo
,
A.
,
Zadrozny
,
T.
,
Sylvester
,
A.
and
Jackson
,
D.
(
2013
)
Fluorescent protein marker lines in maize: generation and applications
.
Int. J. Dev. Biol.
57
,
535
543
59
Chen
,
Z.
,
Zheng
,
W.
,
Chen
,
L.
,
Li
,
C.
,
Liang
,
T.
,
Chen
,
Z.
et al (
2019
)
Green fluorescent protein- and Discosoma sp. red fluorescent protein-tagged organelle marker lines for protein subcellular localization in rice
.
Front. Plant Sci.
10
,
1421
60
Liu
,
Z.
,
Osterlund
,
I.
,
Ruhnow
,
F.
,
Cao
,
Y.
,
Huang
,
G.
,
Cai
,
W.
et al (
2022
)
Fluorescent cytoskeletal markers reveal associations between the actin and microtubule cytoskeleton in rice cells
.
Development
149
,
dev200415
61
Kaduchova
,
K.
,
Marchetti
,
C.
,
Ovecka
,
M.
,
Galuszka
,
P.
,
Bergougnoux
,
V.
,
Samaj
,
J.
et al (
2023
)
Spatial organization and dynamics of chromosomes and microtubules during barley mitosis
.
Plant J.
115
,
602
613
62
Chen
,
T.
,
Wang
,
X.H.
,
von Wangenheim
,
D.
,
Zheng
,
M.Z.
,
Samaj
,
J.
,
Ji
,
W.Q.
et al (
2012
)
Probing and tracking organelles in living plant cells
.
Protoplasma
249
,
157
167
63
Nelson
,
B.K.
,
Cai
,
X.
and
Nebenfuhr
,
A.
(
2007
)
A multicolored set of in vivo organelle markers for co-localization studies in Arabidopsis and other plants
.
Plant J.
51
,
1126
1136
64
Komis
,
G.
,
Novak
,
D.
,
Ovecka
,
M.
,
Samajova
,
O.
and
Samaj
,
J.
(
2018
)
Advances in imaging plant cell dynamics
.
Plant Physiol.
176
,
80
93
65
Reumann
,
S.
,
Quan
,
S.
,
Aung
,
K.
,
Yang
,
P.F.
,
Manandhar-Shrestha
,
K.
,
Holbrook
,
D.
et al (
2009
)
In-depth proteome analysis of Arabidopsis leaf peroxisomes combined with in vivo subcellular targeting verification indicates novel metabolic and regulatory functions of peroxisomes
.
Plant Physiol.
150
,
125
143
66
Schuhmann
,
H.
,
Huesgen
,
P.F.
,
Gietl
,
C.
and
Adamska
,
I.
(
2008
)
The DEG15 serine protease cleaves peroxisomal targeting signal 2-containing proteins in arabidopsis
.
Plant Physiol.
148
,
1847
1856
67
Zhang
,
D.
,
Wadsworth
,
P.
and
Hepler
,
P.K.
(
1993
)
Dynamics of microfilaments are similar, but distinct from microtubules during cytokinesis in living, dividing plant cells
.
Cell Motil. Cytoskel.
24
,
151
155
68
Wang
,
Y.S.
,
Yoo
,
C.M.
and
Blancaflor
,
E.B.
(
2008
)
Improved imaging of actin filaments in transgenic Arabidopsis plants expressing a green fluorescent protein fusion to the C- and N-termini of the fimbrin actin-binding domain 2
.
New Phytol.
177
,
525
536
69
Dyachok
,
J.
,
Sparks
,
J.A.
,
Liao
,
F.
,
Wang
,
Y.S.
and
Blancaflor
,
E.B.
(
2014
)
Fluorescent protein-based reporters of the actin cytoskeleton in living plant cells: fluorophore variant, actin binding domain, and promoter considerations
.
Cytoskeleton (Hoboken)
71
,
311
327
70
Marc
,
J.
,
Granger
,
C.L.
,
Brincat
,
J.
,
Fisher
,
D.D.
,
Kao
,
T.
,
McCubbin
,
A.G.
et al (
1998
)
A GFP-MAP4 reporter gene for visualizing cortical microtubule rearrangements in living epidermal cells
.
Plant Cell
10
,
1927
1940
71
Ben-Nissan
,
G.
,
Cui
,
W.
,
Kim
,
D.J.
,
Yang
,
Y.
,
Yoo
,
B.C.
and
Lee
,
J.Y.
(
2008
)
Arabidopsis casein kinase 1-like 6 contains a microtubule-binding domain and affects the organization of cortical microtubules
.
Plant Physiol.
148
,
1897
1907
72
Ueda
,
K.
,
Matsuyama
,
T.
and
Hashimoto
,
T.
(
1999
)
Visualization of microtubules in living cells of transgenic Arabidopsis thaliana
.
Protoplasma
206
,
201
206
73
Nakamura
,
M.
,
Naoi
,
K.
,
Shoji
,
T.
and
Hashimoto
,
T.
(
2004
)
Low concentrations of propyzamide and oryzalin alter microtubule dynamics in Arabidopsis epidermal cells
.
Plant Cell Physiol.
45
,
1330
1334
74
Liu
,
W.
,
Wang
,
C.
,
Wang
,
G.
,
Ma
,
Y.
,
Tian
,
J.
,
Yu
,
Y.
et al (
2019
)
Towards a better recording of microtubule cytoskeletal spatial organization and dynamics in plant cells
.
J. Integr. Plant Biol.
61
,
388
393
75
Kasahara
,
M.
,
Kagawa
,
T.
,
Oikawa
,
K.
,
Suetsugu
,
N.
,
Miyao
,
M.
and
Wada
,
M.
(
2002
)
Chloroplast avoidance movement reduces photodamage in plants
.
Nature
420
,
829
832
76
Kong
,
S.G.
and
Wada
,
M.
(
2014
)
Recent advances in understanding the molecular mechanism of chloroplast photorelocation movement
.
Biochim. Biophys. Acta
1837
,
522
530
77
Kong
,
S.G.
and
Wada
,
M.
(
2016
)
Molecular basis of chloroplast photorelocation movement
.
J. Plant Res.
129
,
159
166
78
Kong
,
S.G.
and
Wada
,
M.
(
2011
)
New insights into dynamic actin-based chloroplast photorelocation movement
.
Mol. Plant
4
,
771
781
79
Wada
,
M.
(
2016
)
Chloroplast and nuclear photorelocation movements
.
Proc. Jpn Acad. Ser. B Phys. Biol. Sci.
92
,
387
411
80
Gotoh
,
E.
,
Suetsugu
,
N.
,
Yamori
,
W.
,
Ishishita
,
K.
,
Kiyabu
,
R.
,
Fukuda
,
M.
et al (
2018
)
Chloroplast accumulation response enhances leaf photosynthesis and plant biomass production
.
Plant Physiol.
178
,
1358
1369
81
Higa
,
T.
,
Suetsugu
,
N.
,
Kong
,
S.G.
and
Wada
,
M.
(
2014
)
Actin-dependent plastid movement is required for motive force generation in directional nuclear movement in plants
.
Proc. Natl Acad. Sci. U.S.A.
111
,
4327
4331
82
Higa
,
T.
,
Suetsugu
,
N.
and
Wada
,
M.
(
2014
)
Plant nuclear photorelocation movement
.
J. Exp. Bot.
65
,
2873
2881
83
Wang
,
X.L.
,
Gao
,
X.Q.
and
Wang
,
X.C.
(
2011
)
Stochastic dynamics of actin filaments in guard cells regulating chloroplast localization during stomatal movement
.
Plant Cell Environ.
34
,
1248
1257
84
Chu
,
C.P.
,
Liu
,
Z.H.
,
Hu
,
Z.Y.
and
Wang
,
X.L.
(
2011
)
Tubular actin filaments in tobacco guard cells
.
Plant Signal. Behav.
6
,
1578
1580
85
Yuan
,
N.
,
Mendu
,
L.
,
Ghose
,
K.
,
Witte
,
C.S.
,
Frugoli
,
J.
and
Mendu
,
V.
(
2023
)
FKF1 interacts with CHUP1 and regulates chloroplast movement in Arabidopsis
.
Plants (Basel)
12
,
542
86
Aihara
,
Y.
,
Tabata
,
R.
,
Suzuki
,
T.
,
Shimazaki
,
K.
and
Nagatani
,
A.
(
2008
)
Molecular basis of the functional specificities of phototropin 1 and 2
.
Plant J.
56
,
364
375
87
Wada
,
M.
and
Kong
,
S.G.
(
2018
)
Actin-mediated movement of chloroplasts
.
J. Cell Sci.
131
,
jcs210310
88
Paves
,
H.
and
Truve
,
E.
(
2007
)
Myosin inhibitors block accumulation movement of chloroplasts in Arabidopsis thaliana leaf cells
.
Protoplasma
230
,
165
169
89
Suetsugu
,
N.
,
Higa
,
T.
,
Gotoh
,
E.
and
Wada
,
M.
(
2016
)
Light-induced movements of chloroplasts and nuclei are regulated in both Cp-actin-filament-dependent and -independent manners in Arabidopsis thaliana
.
PLoS ONE
11
,
e0157429
90
Labuz
,
J.
,
Hermanowicz
,
P.
and
Gabrys
,
H.
(
2015
)
The impact of temperature on blue light induced chloroplast movements in Arabidopsis thaliana
.
Plant Sci.
239
,
238
249
91
Kitashova
,
A.
,
Schneider
,
K.
,
Furtauer
,
L.
,
Schroder
,
L.
,
Scheibenbogen
,
T.
,
Furtauer
,
S.
et al (
2021
)
Impaired chloroplast positioning affects photosynthetic capacity and regulation of the central carbohydrate metabolism during cold acclimation
.
Photosynth. Res.
147
,
49
60
92
Kodama
,
Y.
,
Tsuboi
,
H.
,
Kagawa
,
T.
and
Wada
,
M.
(
2008
)
Low temperature-induced chloroplast relocation mediated by a blue light receptor, phototropin 2, in fern gametophytes
.
J. Plant Res.
121
,
441
448
93
Ogasawara
,
Y.
,
Ishizaki
,
K.
,
Kohchi
,
T.
and
Kodama
,
Y.
(
2013
)
Cold-induced organelle relocation in the liverwort Marchantia polymorpha L
.
Plant Cell Environ.
36
,
1520
1528
94
Yong
,
L.K.
,
Tsuboyama
,
S.
,
Kitamura
,
R.
,
Kurokura
,
T.
,
Suzuki
,
T.
and
Kodama
,
Y.
(
2021
)
Chloroplast relocation movement in the liverwort Apopellia endiviifolia
.
Physiol. Plant
173
,
775
787
95
Fujii
,
Y.
,
Tanaka
,
H.
,
Konno
,
N.
,
Ogasawara
,
Y.
,
Hamashima
,
N.
,
Tamura
,
S.
et al (
2017
)
Phototropin perceives temperature based on the lifetime of its photoactivated state
.
Proc. Natl Acad. Sci. U.S.A.
114
,
9206
9211
96
Hirano
,
S.
,
Sasaki
,
K.
,
Osaki
,
Y.
,
Tahara
,
K.
,
Takahashi
,
H.
,
Takemiya
,
A.
et al (
2022
)
The localization of phototropin to the plasma membrane defines a cold-sensing compartment in Marchantia polymorpha
.
PNAS Nexus
1
,
pgac030
97
Sakata
,
M.
,
Kimura
,
S.
,
Fujii
,
Y.
,
Sakai
,
T.
and
Kodama
,
Y.
(
2019
)
Relationship between relocation of phototropin to the chloroplast periphery and the initiation of chloroplast movement in Marchantia polymorpha
.
Plant Direct
3
,
e00160
98
Fujii
,
Y.
,
Ogasawara
,
Y.
,
Takahashi
,
Y.
,
Sakata
,
M.
,
Noguchi
,
M.
,
Tamura
,
S.
et al (
2020
)
The cold-induced switch in direction of chloroplast relocation occurs independently of changes in endogenous phototropin levels
.
PLoS ONE
15
,
e0233302
99
Kimura
,
S.
and
Kodama
,
Y.
(
2016
)
Actin-dependence of the chloroplast cold positioning response in the liverwort Marchantia polymorpha L
.
PeerJ
4
,
e2513
100
Yong
,
L.K.
and
Kodama
,
Y.
(
2023
)
Dark-induced chloroplast relocation depends on actin filaments in the liverwort Apopellia endiviifolia along with the light- and cold-induced relocations
.
Plant Cell Environ.
46
,
1822
1832
101
Tanaka
,
H.
,
Sato
,
M.
,
Ogasawara
,
Y.
,
Hamashima
,
N.
,
Buchner
,
O.
,
Holzinger
,
A.
et al (
2017
)
Chloroplast aggregation during the cold-positioning response in the liverwort Marchantia polymorpha
.
J. Plant Res.
130
,
1061
1070
102
Ovsyannikov
,
A.Y.
and
Koteyeva
,
N.K.
(
2020
)
Seasonal movement of chloroplasts in mesophyll cells of two Picea species
.
Protoplasma
257
,
183
195
103
Kumar
,
A.S.
,
Park
,
E.
,
Nedo
,
A.
,
Alqarni
,
A.
,
Ren
,
L.
,
Hoban
,
K.
et al (
2018
)
Stromule extension along microtubules coordinated with actin-mediated anchoring guides perinuclear chloroplast movement during innate immunity
.
eLife
7
,
e23625
104
Ding
,
X.
,
Jimenez-Gongora
,
T.
,
Krenz
,
B.
and
Lozano-Duran
,
R.
(
2019
)
Chloroplast clustering around the nucleus is a general response to pathogen perception in Nicotiana benthamiana
.
Mol. Plant Pathol.
20
,
1298
1306
105
Park
,
E.
,
Caplan
,
J.L.
and
Dinesh-Kumar
,
S.P.
(
2018
)
Dynamic coordination of plastid morphological change by cytoskeleton for chloroplast-nucleus communication during plant immune responses
.
Plant Signal. Behav.
13
,
e1500064
106
Caplan
,
J.L.
,
Kumar
,
A.S.
,
Park
,
E.
,
Padmanabhan
,
M.S.
,
Hoban
,
K.
,
Modla
,
S.
et al (
2015
)
Chloroplast stromules function during innate immunity
.
Dev. Cell
34
,
45
57
107
Erickson
,
J.L.
,
Kantek
,
M.
and
Schattat
,
M.H.
(
2017
)
Plastid-nucleus distance alters the behavior of stromules
.
Front. Plant Sci.
8
,
1135
108
Savage
,
Z.
,
Duggan
,
C.
,
Toufexi
,
A.
,
Pandey
,
P.
,
Liang
,
Y.
,
Segretin
,
M.E.
et al (
2021
)
Chloroplasts alter their morphology and accumulate at the pathogen interface during infection by Phytophthora infestans
.
Plant J.
107
,
1771
1787
109
Irieda
,
H.
and
Takano
,
Y.
(
2021
)
Epidermal chloroplasts are defense-related motile organelles equipped with plant immune components
.
Nat. Commun.
12
,
2739
110
Irieda
,
H.
(
2022
)
Emerging roles of motile epidermal chloroplasts in plant immunity
.
Int. J. Mol. Sci.
23
,
4043
111
Hanson
,
M.R.
and
Conklin
,
P.L.
(
2020
)
Stromules, functional extensions of plastids within the plant cell
.
Curr. Opin. Plant Biol.
58
,
25
32
112
Fujiwara
,
M.T.
,
Yoshioka
,
Y.
,
Hirano
,
T.
,
Kazama
,
Y.
,
Abe
,
T.
,
Hayashi
,
K.
et al (
2012
)
Visualization of plastid movement in the pollen tube of Arabidopsis thaliana
.
Plant Signal. Behav.
7
,
34
37
113
Fujiwara
,
M.T.
,
Hashimoto
,
H.
,
Kazama
,
Y.
,
Hirano
,
T.
,
Yoshioka
,
Y.
,
Aoki
,
S.
et al (
2010
)
Dynamic morphologies of pollen plastids visualised by vegetative-specific FtsZ1-GFP in Arabidopsis thaliana
.
Protoplasma
242
,
19
33
114
Logan
,
D.C.
and
Leaver
,
C.J.
(
2000
)
Mitochondria-targeted GFP highlights the heterogeneity of mitochondrial shape, size and movement within living plant cells
.
J. Exp. Bot.
51
,
865
871
PMID:
[PubMed]
115
Zhang
,
Y.
,
Sheng
,
X.
,
Meng
,
X.
and
Li
,
Y.
(
2014
)
The circular F-actin bundles provide a track for turnaround and bidirectional movement of mitochondria in Arabidopsis root hair
.
PLoS ONE
9
,
e91501
116
Zheng
,
M.
,
Wang
,
Q.
,
Teng
,
Y.
,
Wang
,
X.
,
Wang
,
F.
,
Chen
,
T.
et al (
2010
)
The speed of mitochondrial movement is regulated by the cytoskeleton and myosin in Picea wilsonii pollen tubes
.
Planta
231
,
779
791
117
Van Gestel
,
K.
,
Kohler
,
R.H.
and
Verbelen
,
J.P.
(
2002
)
Plant mitochondria move on F-actin, but their positioning in the cortical cytoplasm depends on both F-actin and microtubules
.
J. Exp. Bot.
53
,
659
667
118
Lovy-Wheeler
,
A.
,
Cardenas
,
L.
,
Kunkel
,
J.G.
and
Hepler
,
P.K.
(
2007
)
Differential organelle movement on the actin cytoskeleton in lily pollen tubes
.
Cell Motil. Cytoskeleton
64
,
217
232
119
Colaco
,
R.
,
Moreno
,
N.
and
Feijo
,
J.A.
(
2012
)
On the fast lane: mitochondria structure, dynamics and function in growing pollen tubes
.
J. Microsc.
247
,
106
118
120
Zhang
,
S.
,
Wang
,
C.
,
Xie
,
M.
,
Liu
,
J.
,
Kong
,
Z.
and
Su
,
H.
(
2018
)
Actin bundles in the pollen tube
.
Int. J. Mol. Sci.
19
,
3710
121
Tian
,
X.
,
Wang
,
X.
and
Li
,
Y.
(
2021
)
Myosin XI-B is involved in the transport of vesicles and organelles in pollen tubes of Arabidopsis thaliana
.
Plant J.
108
,
1145
1161
122
Romagnoli
,
S.
,
Cai
,
G.
,
Faleri
,
C.
,
Yokota
,
E.
,
Shimmen
,
T.
and
Cresti
,
M.
(
2007
)
Microtubule- and actin filament-dependent motors are distributed on pollen tube mitochondria and contribute differently to their movement
.
Plant Cell Physiol.
48
,
345
361
123
Ni
,
C.Z.
,
Wang
,
H.Q.
,
Xu
,
T.
,
Qu
,
Z.
and
Liu
,
G.Q.
(
2005
)
AtKP1, a kinesin-like protein, mainly localizes to mitochondria in Arabidopsis thaliana
.
Cell Res.
15
,
725
733
124
Wang
,
X.
,
Sheng
,
X.
,
Tian
,
X.
,
Zhang
,
Y.
and
Li
,
Y.
(
2020
)
Organelle movement and apical accumulation of secretory vesicles in pollen tubes of Arabidopsis thaliana depend on class XI myosins
.
Plant J.
104
,
1685
1697
125
Zhang
,
L.
and
Xing
,
D.
(
2008
)
Methyl jasmonate induces production of reactive oxygen species and alterations in mitochondrial dynamics that precede photosynthetic dysfunction and subsequent cell death
.
Plant Cell Physiol.
49
,
1092
1111
126
Zhang
,
L.
,
Li
,
Y.
,
Xing
,
D.
and
Gao
,
C.
(
2009
)
Characterization of mitochondrial dynamics and subcellular localization of ROS reveal that HsfA2 alleviates oxidative damage caused by heat stress in Arabidopsis
.
J. Exp. Bot.
60
,
2073
2091
127
Gao
,
C.
,
Xing
,
D.
,
Li
,
L.
and
Zhang
,
L.
(
2008
)
Implication of reactive oxygen species and mitochondrial dysfunction in the early stages of plant programmed cell death induced by ultraviolet-C overexposure
.
Planta
227
,
755
767
128
Islam
,
M.S.
,
Niwa
,
Y.
and
Takagi
,
S.
(
2009
)
Light-dependent intracellular positioning of mitochondria in Arabidopsis thaliana mesophyll cells
.
Plant Cell Physiol.
50
,
1032
1040
129
Oikawa
,
K.
,
Imai
,
T.
,
Thagun
,
C.
,
Toyooka
,
K.
,
Yoshizumi
,
T.
,
Ishikawa
,
K.
et al (
2021
)
Mitochondrial movement during its association with chloroplasts in Arabidopsis thaliana
.
Commun. Biol.
4
,
292
130
Rai
,
A.K.
,
Chen
,
T.
and
Moroney
,
J.V.
(
2021
)
Mitochondrial carbonic anhydrases are needed for optimal photosynthesis at low CO2 levels in Chlamydomonas
.
Plant Physiol.
187
,
1387
1398
131
Fu
,
X.
and
Walker
,
B.J.
(
2023
)
Dynamic response of photorespiration in fluctuating light environments
.
J. Exp. Bot.
74
,
600
611
132
Hu
,
J.
and
Desai
,
M.
(
2008
)
Light control of peroxisome proliferation during Arabidopsis photomorphogenesis
.
Plant Signal. Behav.
3
,
801
803
133
Desai
,
M.
and
Hu
,
J.
(
2008
)
Light induces peroxisome proliferation in Arabidopsis seedlings through the photoreceptor phytochrome A, the transcription factor HY5 HOMOLOG, and the peroxisomal protein PEROXIN11b
.
Plant Physiol.
146
,
1117
1127
134
Desai
,
M.
,
Pan
,
R.
and
Hu
,
J.
(
2017
)
Arabidopsis forkhead-associated domain protein 3 negatively regulates peroxisome division
.
J. Integr. Plant Biol.
59
,
454
458
135
Piacentini
,
D.
,
Corpas
,
F.J.
,
D'Angeli
,
S.
,
Altamura
,
M.M.
and
Falasca
,
G.
(
2020
)
Cadmium and arsenic-induced-stress differentially modulates arabidopsis root architecture, peroxisome distribution, enzymatic activities and their nitric oxide content
.
Plant Physiol. Biochem.
148
,
312
323
136
Rodriguez-Serrano
,
M.
,
Romero-Puertas
,
M.C.
,
Sanz-Fernandez
,
M.
,
Hu
,
J.
and
Sandalio
,
L.M.
(
2016
)
Peroxisomes extend peroxules in a fast response to stress via a reactive oxygen species-mediated induction of the peroxin PEX11a
.
Plant Physiol.
171
,
1665
1674
137
Terron-Camero
,
L.C.
,
Rodriguez-Serrano
,
M.
,
Sandalio
,
L.M.
and
Romero-Puertas
,
M.C.
(
2020
)
Nitric oxide is essential for cadmium-induced peroxule formation and peroxisome proliferation
.
Plant Cell Environ.
43
,
2492
2507
138
Jedd
,
G.
and
Chua
,
N.H.
(
2002
)
Visualization of peroxisomes in living plant cells reveals acto-myosin-dependent cytoplasmic streaming and peroxisome budding
.
Plant Cell Physiol.
43
,
384
392
139
Peremyslov
,
V.V.
,
Prokhnevsky
,
A.I.
,
Avisar
,
D.
and
Dolja
,
V.V.
(
2008
)
Two class XI myosins function in organelle trafficking and root hair development in Arabidopsis
.
Plant Physiol.
146
,
1109
1116
140
Barton
,
K.
,
Mathur
,
N.
and
Mathur
,
J.
(
2013
)
Simultaneous live-imaging of peroxisomes and the ER in plant cells suggests contiguity but no luminal continuity between the two organelles
.
Front. Physiol.
4
,
196
141
Rodriguez-Serrano
,
M.
,
Romero-Puertas
,
M.C.
,
Sparkes
,
I.
,
Hawes
,
C.
,
del Rio
,
L.A.
and
Sandalio
,
L.M.
(
2009
)
Peroxisome dynamics in arabidopsis plants under oxidative stress induced by cadmium
.
Free Radic. Biol. Med.
47
,
1632
1639
142
Gao
,
H.
,
Metz
,
J.
,
Teanby
,
N.A.
,
Ward
,
A.D.
,
Botchway
,
S.W.
,
Coles
,
B.
et al (
2016
)
In vivo quantification of peroxisome tethering to chloroplasts in tobacco epidermal cells using optical tweezers
.
Plant Physiol.
170
,
263
272
This is an open access article published by Portland Press Limited on behalf of the Biochemical Society and distributed under the Creative Commons Attribution License 4.0 (CC BY-NC-ND).

Supplementary data