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Commentary
Polar auxin transport in relation to
long-distance transport of nutrients
in the Charales
John A. Raven*,†
Division of Plant Sciences, University of Dundee at the James
Hutton Institute, Invergowrie, Dundee DD2 5DA, UK (permanent
address) and Plant Functional Biology and Climate Change
Cluster, University of Technology Sydney, PO Box 123, NSW
2007, Australia
* To whom correspondence should be addressed. E-mail:
[email protected]
Journal of Experimental Botany Vol. 64, No. 1, pp. 1–9, 2013
doi:10.1093/jxb/ers358
Abstract
This paper examines the significance of the recent demonstration of polar auxin transport (PAT) in the green macroalga Chara (Charophyceae: Charales) and, especially,
options for explaining some features of PAT in the Charales.
The occurrence of PAT in the Charales shows that PAT
originated in the algal ancestors of the embryophytes
(liverworts, mosses, hornworts, and vascular plants),
although it is not yet known if PAT occurs elsewhere in the
Charophyceae or in other algae. While in the embryophytes
PAT occurs in parenchymatously constructed structures
which commonly also have xylem and phloem (or their bryophyte analogues) as long-distance transport processes in
parallel to PAT, in Chara corallina PAT shares the pathway
for long-distance transport of nutrients though the parenchymatously constructed nodal complexes and the single
giant cells of the internode. The speed of auxin movement
of PAT is much more rapid than that attributable to diffusion
and of the same order as the rate of cytoplasmic streaming in the giant internodal cells, yet complete inhibition of
streaming by the inhibitor cytochalasin H does not slow
down auxin transport. Explanations for this phenomenon
are sought in the operation of other mechanochemical
motors, dynein–tubulin and kinesin–tubulin, as alternatives to the myosin–actin system which powers cytoplasmic streaming. Experiments in which microtubules are
disrupted, for example by colchicine, could show if one
of the tubulin-based motors is involved. If these motors
are involved, some mechanism is needed to amplify the
speeds known for the motors to explain the order of magnitude higher speeds seen for auxin transport.
Key words: Chara, Charophyceae, colchicine, cytochalasin,
cytoplasmic streaming, Nitella, nutrient transport, polar auxin
transport, rhizoids.
Introduction
Darwin and Darwin (1880) performed experiments on flowering plant tropisms from which they inferred the occurrence and differential concentrations of an endogenous
compound, now known to be indoleacetic acid (auxin) (see
Ayres, 2008; Leyser, 2010). The functioning of auxin relates
in part to its polar transport and, for tropisms, to its lateral
transport. A widely accepted mechanism for polar auxin
transport (PAT) is the chemiosmotic hypothesis (Rubery
and Sheldrake, 1973, 1974; Raven, 1975; Goldsmith, 1977;
Hošek et al., 2012). PAT has been identified in the sporophyte phase of vascular plants and mosses (Fujita et al.,
2008); Boot et al. (2012) have now shown that PAT also
occurs in the Charales, as exemplified by Chara corallina
Klein ex C.J. Wildenow and Chara vulgaris var. longibracteata (Kützing) J Grives & Bullock-Webster. The significance
of this work is that the Charales are an order of the algal
division Charophyta which, with the Embryophyta (extant
liverworts, hornworts, mosses, and vascular plants, and
many extinct taxa), comprise the Streptobionta, with the
Charales among the closest living relatives of the ancestors
of the embryophytes (Graham et al., 2009; Taylor et al.,
2009; Raven and Edwards, 2013). As Boot et al. (2012)
point out, the Charales have a morphology superficially
resembling that of the sporophytes of vascular plants,
although the ‘plant’ of the Charales is, in life cycle terms,
the equivalent of the gametophyte phase of the embryophyte life cycle. Furthermore, the shoot of the Charales is
made up of giant cylindrical internodal cells (with or without corticating cells) separated by nodes which bear whorls
of ‘leaves’, again composed of giant cells, and branches.
Growth is apical.
In this paper we consider several aspects of the findings
of Boot et al. (2012). Special attention is given to the phylogenetic occurrence of PAT, the functioning of rhizoids in
relation to PAT, and particularly to explaining the speed of
auxin movement in PAT in the context of the cytochalasin
insensitivity of PAT at concentrations of cytochalasin which
completely inhibit cytoplasmic streaming.
Phylogenetic considerations of polar auxin
transport in the Charales
Plants of the Charales are haploid, although the giant internodal cells are multinucleate and/or have several large nuclei
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2 | Raven et al.
with many copies of the haploid set of chromosomes (Shen,
1967). Our present, incomplete, understanding suggests that
PAT in embryophytes is restricted to the sporophyte phase
and does not occur in free-living gametophyte phases of
non-seed plants (Fujita et al., 2008; Boot et al., 2012). This is
of interest because the morphology of the gametophytes of
many mosses resembles that of vascular plant sporophytes,
though with very different anatomy (e.g. no intercellular gas
spaces or stomata, hydrome rather than xylem, and leptome
rather than phloem). Long-distance nutrient transport in
the Charales, apparently involving cytoplasmic streaming
(Raven, 2003), is the functional equivalent of nutrient transport in the xylem and phloem in the parenchymatously constructed sporophytes of vascular embryophytes. Apoplasmic
xylem transport, driven by transpiration and root pressure,
and symplasmic phloem transport, using the Münch mechanism (Pickard, 2012), comprise the long-distance water
nutrient transport pathways of terrestrial vascular plant
sporophytes. Phloem transport occurs in secondarily submerged aquatic vascular plants, as can xylem transport, at
least to the extent that xylem flow driven by root pressure
occurs (Raven, 1984). PAT in the vascular plants and moss
sporophytes occurs in parenchymatous tissue independent of the xylem and phloem. Xylem-like (hydrome) and
phloem-like (leptome) also occur in some moss sporophytes,
as well as in some moss gametophytes (Raven, 2003), and
again PAT occurs in sporophytes through files of parenchyma cells. In contrast, PAT in the Charales involves the
same cells as are involved in axial nutrient transport; the
internodal cells in this pathway show rapid cytoplasmic
streaming. Further analysis of PAT and its relationship to
long-distance nutrient transport, and the role of rhizoids, in
the Characeae is considered in detail after brief discussions
of the phylogeny of PAT and the relationship of PAT to
rhizoids of the Charales.
Phylogeny of polar auxin transport
There are several independent lines of evolution of multicellular (not just colonial: Beardall et al., 2009) algae with varying degrees of differentiation (Graham et al., 2009). Among
the green algae, as well as the Charophyta (as exemplified by
the Charales which are probably the closest living relatives
of the ancestors of the embryophytes), there are variously
differentiated multicellular members of the Chlorophyceae
(Draparnaldia, Fritschiella), Trebouxiophyceae (Prasiola).
and Ulvophyceae (Trentepholia) (Graham et al., 2009; Raven
and Edwards, 2013). In the Rhodophyta, multicellularity
evolved in both the Bangiophyceae (e.g. Bangiales) and the
Floridiophyceae (all species), while in the Heterokontophyta
all of the species of the Fucophyceae (= Phaeophyceae) are
multicellular. The involvement of auxin, and its (polar?)
transport in the morphogenesis of these multicellular algae
is still uncertain, for example the work on the Fucophyceae
of Sun et al. (2004) and Le Bail et al. (2010). The best prospects for rapid progress probably occur for the fucophycean
Ectocarpus siliculosus (Dillwyn) Lyngbe whose genome
has been fully sequences and annotated (Sun et al., 2010).
If PAT of the kind found in the Streptobionta occurs in
algal clades other than the Charophyceae, it would presumably have arisen from horizontal gene transfer, since
the Charophyceae and other multicellular green algae, and
non-green multicellular algae, all arose independently from
unicellular ancestors (Graham et al., 2009). There would be
no opportunity for PAT in the differentiated but unicellular
(otherwise known as acellular) macrophytic members of the
Ulvophyceae such as Caulerpa (Raven, 1981, 1984; Graham
et al., 2009).
The absence of genes encoding auxin influx carriers in the
charophyceans Coleochaete and Spirogyra is discussed by
Boot et al. (2012) in terms of the role of the carriers in recovering leaked auxin and preventing leakage to surrounding
cells (Reinhardt. 2003); this is part of their proposed role in
the symplasm–apoplasm–symplasm transfer which underlies
PAT (fig. 1 of Blakeslee et al., 2005; Kramer et al., 2006).
Boot et al. (2012) suggest that an auxin influx carrier may not
be involved in PAT in the characeans, based on observations
of auxin efflux. While all tissues in which PAT has been characterized (embryophyte sporophytes; Chara) or may occur
(some complex brown algae: Sun et al., 2004; Le Bail et al.,
2010) have plasmodesmata, PAT does not involve plasmodesmata. Movement of auxin into the phloem, where transport
is by mass flow and is not part of the PAT process, also seems
to involve the auxin influx transporters (Blakeslee et al., 2005;
Hošek et al., 2012).
Rhizoids of Chara in relation to polar auxin
transport
The Charales are rhizophytes (i.e. they are attached to a
substrate with small particle size relative to the size of the
attachment organs; Raven, 1981, 1984). In the case of the
Charales, attachment is via rhizoids which serve in nutrient
uptake from the sediment, discussed below in the context
of axial nutrient transport. In some cases, mature stands of
characeans, specifically Chara hispida var. major (Hartm.)
RD Wood, produce very few rhizoids and so are not attached
to the substratum but are supported by their neighbours and
rely largely on internal recycling of nutrients such as nitrogen and phosphorus, with apical growth and basal decay
(Andrews et al., 1984a). Both decay and growth are faster in
the summer, so the biomass is essentially constant throughout the year in this perennial organism (Andrews et al.,
1984a). However, rhizoids are needed for both attachment
and nutrient uptake in the establishment and net growth
phases of the Charales.
Rhizoids of the Charales are multicellular; in this they differ from the rhizoids of the free-living gametophytes and the
root hairs of the free-living sporophytes of embryophytes.
The characean rhizoids are unbranched, resembling embryophyte rhizoids and root hairs, but differing from the branched
mycorrhizal fungi (Smith and Read, 2008). Rhizoid cells of
the Charales, like the shoot cells, have cytoplasmic streaming
(Ackers et al., 2000). The multicellularity of the characean
Commentary | 3
rhizoid only extends to the occurrence of cross-walls rather
than the parenchymatous structure of the nodes of characean shoots or the cortication of the giant internodal cells
of some species of Chara, for example the corticate C. vulgaris var. longibracteata used by Boot et al. (2012) for comparison with the ecorticate C. corallina. The rhizoids of the
characeans grow apically, as, for mechanical reasons, do all
plant or algal organs growing through soil or sediments, or
growing over solid substrata (Raven and Edwards, 2001). The
growth of characean rhizoids requires, or is at least stimulated by, auxin: while the growth of rhizoids of intact algae
is not influenced by exogenously supplied auxin, decapitated
(shoot removed) algae show significant stimulation of rhizoid
growth by exogenous auxin (Klämbt et al., 1992). An inhibitor (α-naphthylphthalamic acid) of the efflux of the auxin
anion through the PIN transporters inhibits rhizoid growth
in intact plants, and increases the retention of labelled auxin
by explants, although it not clear whether these effects are
direct effects on the rhizoids or a consequence of inhibition
of PAT down the shoot (Klämbt et al., 1992).
Charalean rhizoids are gravitropic, using BaSO4 crystals as
the particles which move relative to the rest of the cell (Raven
and Knoll, 2010). In roots of vascular plants, the cells which
execute the tropic response to the gravitational stimulus are
not the cells which perceive the stimulus, and lateral (polar)
auxin transport is a component of the signalling chain. Does
a similar system operate in characean rhizoids? While there is
no direct evidence, this seems unlikely because in the Charales
the BaSO4 particles are in the same cell (and the same part
of the cell) in which the tropic differential growth occurs.
Gravisensing seems to involve contact of the statoliths with
membrane-bound receptor molecules (Limbach et al., 2005).
In any case, the maintenance of a lateral gradient of auxin
across a cell of the diameter of the rhizoid would be next to
impossible (see Nobel, 2009).
The role of cytoplasmic streaming in
the polar transport of auxin and the
axial transport of nutrient solutes in the
Charales
Absence of inhibition of polar auxin transport by
cytochalasin H at concentrations which inhibit
cytoplasmic streaming
Boot et al. (2012) showed that the speed of PAT in Chara
was much greater than could be accounted for by diffusive
intracellular movement and was of a similar magnitude to the
speed of cytoplasmic streaming. The minimum estimate of the
speed of auxin transport along C. corallina axes of 11–14 µm
s–1 are about a third of the measured speed of cytoplasmic
streaming in the same cells, namely 33–56 µm s–1 (Boot et al.,
2012), which is similar to values of 50–60 µm s–1, and sometimes up to 100 µm s–1 in C. corallina and other characeans at
20–25 °C (Hope and Walker, 1975; Raven and Smith, 1978,
Pickard, 2003; Raven, 2003; Shimmen and Yokota, 2004;
Verchot-Lubicz and Goldman, 2010). However, Boot et al.
(2012) point out that their measurements of the speed of
auxin transport are likely to be underestimates. While there is
substantial agreement among different methods on the speed
of cytoplasmic streaming (Pickard, 2003; Verchot-Lubicz
and Goldman, 2010; van de Meent et al., 2010), it is difficult
to measure the speed of transport or movement of a solute
within plants (Pickard, 2003; Boot et al., 2012)
Despite cytoplasmic streaming in giant internodal cells
being the obvious explanation for the speed of PAT in
Chara, PAT was not inhibited by cytochalasin H at concentrations which completely inhibited cytoplasmic streaming. There is some inhibition of PAT in flowering plants by
cytochalasin at concentrations which completely inhibit
cytoplasmic streaming (Cande et al., 1973; Butler et al.,
1998; Muday et al., 2000). However, it is possible that this
inhibition relates to the dependence on actin filaments of
the production and maintenance of the polar arrangement
of auxin efflux carriers rather than to the intracellular flux
of auxin (Butler et al., 1998; Muday et al., 2000). To put
the independence of very rapid (by comparison with diffusion) PAT in Chara from cytoplasmic streaming into context, the evidence on the axial transport of nutrients is now
considered.
Cytoplasmic streaming in the long-distance (axial)
transport of nutrient solutes in the Charales
Axial transport of nutrients in characeans is commonly
thought to involve cytoplasmic streaming for intracellular
movement and diffusion through plasmodesmata for intercellular transport (Hope and Walker, 1975; Bostrom and
Walker, 1976; Raven, 1981; Ding and Tazawa, 1989; Raven,
2003). Bostrom and Walker (1976) studied the movement
of 36Cl– between adjacent cells of C. corallina as influenced
by the rate of cytoplasmic streaming which was varied by
the addition of a range of concentrations of the inhibitor
cytochalasin B. Their work showed that cytochalasin B had
no effect on the active influx of 36Cl– into the internodal cells,
but decreased the transfer of 36Cl– from the cell to which the
isotope had been supplied into the adjacent internodal cell in
direct proportion to the decrease in the speed of cytoplasmic
streaming. Bostrom and Walker (1976) used the most economical hypothesis to explain these data, namely that the
rate at which 36Cl– arrives at the plasmodesmata linking the
nodal cell to the nodal complex of small cells in parenchymatous tissue determines the rate of symplasmic diffusion of
the anion through the nodal complex, and that this rate of
delivery is proportional to the speed of cytoplasmic streaming. A less economical hypothesis considered by Bostrom and
Walker (1976) is that the cytochalasin effect is on transport
through the nodal complex.
The approach of Bostrom and Walker was followed and
developed further by Ding and Tazawa (1989), again using
C. corallina, although with 86Rb+ (as an analogue of K+) as
the solute, and with temperature as well as cytochalasin B as
the means of decreasing the speed of cytoplasmic streaming.
The results are closely similar to those of Bostrom and Walker
(1976); that is, direct proportionality between transnodal
4 | Raven et al.
solute flux and the speed of cytoplasmic streaming. The
studies of Bostrom and Walker (1976) and Ding and Tazawa
(1989) are apparently the only data sets in which changing
the speed of cytoplasmic streaming was used to examine its
involvement in axial solute transport in the Charales.
Other work consistent with a role for cytoplasmic streaming in axial solute transport in the Charales did not use
cytochalasin, but was still plausibly interpreted in terms of
an essential role for cytoplasmic streaming. Raven and Smith
(1978) and Raven (1981) calculated the differences in the concentration of sucrose between the upward and downward
streams of the streaming cytoplasm of C. corallina which
would be required to account for the organic carbon supply
needed for apical growth of shoots. The calculations involved
determining the requirement for organic carbon for apical
growth of the shoot from the organic carbon content of the
length of shoot added each day. From this is subtracted the
quantity of organic carbon added by photosynthesis minus
respiration over a 24 h light–dark cycle. This calculation
leaves the shortfall of organic carbon which is supplied by
cytoplasmic streaming. The other input is the axial speed of
cytoplasmic streaming, which is less than the measured speed
of streaming, since cytoplasmic streaming follows a spiral
path (Goldstein et al., 2008).
The method equated the difference in sucrose concentration between upward and downward streams expressed
as organic carbon (mol organic carbon in sucrose m–3) to
account for the observed growth rate expressed on the basis
of the cross-sectional area of the upward stream (mol organic
carbon m–2 cross-sectional area s–1) divided by the axial speed
of cytoplasmic streaming (m s–1). The calculated difference
in sucrose concentration between the upward and downward
stream can be readily accommodated by the measured sucrose
concentration in the streaming cytoplasm (Ding et al., 1991),
as discussed by Raven (2003).
Raven (2003) cites work (Andrews et al., 1984b; Box et al.,
1984; Box, 1986, 1987, 1988) showing that phosphorus, and
combined nitrogen, entering the rhizoids of C. hispida move
to the shoots, and their axial movement through the shoot
is consistent with involvement of cytoplasmic streaming.
However, it is important to re-emphasize that neither this
work nor that discussed above on organic carbon flux to the
shoot apex examined the effect of inhibition of cytoplasmic
streaming by cytochalasin on nutrient movement and distribution. The same is true of the work of Vemeer et al. (2003)
on the uptake and distribution of ammonium and nitrate
taken up by rhizoids and shoots of Chara sp., and also the
work of Wüstenberg et al. (2011) showing that C. hispida (L.)
Hartm. grew rapidly when phosphorus was supplied as tricalcium phosphate to the rhizoids in the sediment with no
phosphate in the bulk water round the shoot. There was no
treatment with measurements of growth rate with phosphate
supplied to both rhizoid and shoot; it is clear that internodal
cells of characeans can take up phosphate from the surrounding water (Smith, 1966; Mimura et al., 1998; Reid et al., 2000).
The findings of Boot et al. (2012) on PAT clearly show that
transport of auxin, which involves apoplasmic fluxes for intercellular transport, does not show the direct proportionality
between axial flux and the speed of cytoplasmic streaming
found for symplasmic transport of 36Cl– and of 86Rb+ across
the nodes of Chara. This is despite the clear quantitative
inadequacy of diffusion to explain the observed axial auxin
movement for the segment of the pathway involving movement within internodal cells. It is of interest that Bostrom and
Walker (1976) considered as an explanation for their results,
but rejected as less economical, the hypothesis that the
cytochalasin effect on intercellular transport of 36Cl– is in the
symplasmic flux through the nodal complex rather than the
effect on cytoplasmic streaming. Since there is no symplasmic
step in PAT, the alternative hypothesis would bring the transport of nutrient solutes into line with the PAT. This alternative hypothesis finds some support from the occurrence
of actin and myosin within the plasmodesmatal channel, at
least in flowering plants (White and Barton, 2011). However,
the alternative hypothesis requires the use of a cytochalasininsensitive mechanochemical motor for the intracellular
transport of all of the nutrient solutes.
Possible involvement of mechanochemical motors
other than myosin–actin in polar auxin transport
A further possibility, discussed below, is that auxin transport
could be accounted for as cargo on dynein–tubulin or kinesin–
tubulin mechanochemical motors, either by binding to a protein or in vacuoles undetectable by the light microscopy used
by Boot et al. (2012), with unknown loading and unloading
steps. However, such a mechanism would be impossible for
the major osmotically active solutes in the cytosol, Cl– (tracer
36
Cl–) and K+ (tracer 86Rb+), which are present at concentrations several orders of magnitude greater than that of auxin
(Vorobiev, 1967; Reeves et al., 1985; Petersson et al., 2009).
Assuming that all of the auxin measured in protoplasts of
Arabidopsis thaliana (L.) Heynh.by Petersson et al. (2009) is
in the cytoplasm, and that the cytoplasm occupies 10% of
the protoplast, the auxin concentration in the cytoplasm
(cytosol plus membrane-bounded compartments) is up to
10 µM, while the Cl– concentration is at least 20 mM and the
K+ concentration is ~100 mM (Bradley and Williams, 1967;
Vorobiev, 1967; Larkum, 1969; Lefebvre and Gillet, 1971;
Lüttge and Higinbotham, 1979; Reeves et al., 1985).
The myosin–actin motor which drives cytoplasmic streaming in the Characeae is exceptional in the speed of movement
of the ATPase component (in this case the characean form of
myosin XI) along the skeletal filament (in this case filamentous actin) relative to other myosin–actin systems, and also
dynein–tubulin and kinesin–tubulin motors (Table 1). These
speeds of 60 µm s–1 at 20–25 °C for cytoplamsic streaming
in large internodal cells of characeans in vivo have also been
demonstrated in vitro (Yamamoto et al., 1994; HigashiFujime et al., 1995; Rivolta et al., 1995). The speed is the
product of the step size (i.e. the distance of relative movement of myosin and actin in each ATP hydrolysis cycle) and
the specific rate (s–1) of ATP hydrolysis (i.e. ATP hydrolysis
s–1 per myosin head activated by attachment to filaments at
saturating ATP and filament concentrations). With a step
length (sliding distance per ATP hydrolysed) of 0.019 µm
Commentary | 5
Table 1. Speed of movement and maximum rates of ATP hydrolysis of biological mechanochemical motors. Based on Howard (1997).
Motor
Speed
Speed
ATPase specific References
in vivo µm s–1 in vitro µm s–1 activitya
Myosins of amoeboid
movement, vesicle transport,
smooth and striated muscle
Myosin XI used in streaming of
Characeae
0.2–6
0.2–8
1.2–20
Howard (1997)
50–60 (100)
60
390–500
Dynein in axoneme
Dynein in cytoplasm
Kinesin in mitosis and meiosis
Kinesin in other transport
processes
7
11
0.018
1.8–2.0
4.5
1.25
0.06
0.4–0.8
10
2
2
44
Hope and Walker (1975), Yamamoto et al. (1994, 2006), Higashi-Fujime
et al. (1995), Rivolta et al. (1995), Howard (1997), Ito et al. (2003), Kimura
et al. (2003), Shimmen and Yokota (2004), Ito et al. (2007, 2010)
Howard (1997)
Howard (1997)
Howard (1997)
Howard (1997)
a
ATP hydrolysis s–1 per myosin/dynein/kinesin head activated by attachment to actin (for myosin) or tubulin (for dynein and kinesin) filaments
at saturating ATP and filament concentrations.
(Kimura et al., 2003) and a specific ATP hydrolysis rate of
500 s–1 (Ito et al., 2003, 2007), the predicted speed of movement is 9.5 µm s–1, which is still only 16% of the observed
speed in vitro and in vivo. Kimura et al. (2003) suggest that
this discrepancy could be explained if the ATP hydrolysis
rate increases if multiple myosin molecules are moving along
the same actin filament.
As already indicated, the alternative cytochalasin-insensitive mechanochemical motors (dynein–tubulin or kinesin–
tubulin) have sliding speeds (Table 1) which are inadequate
by at least an order of magnitude to account for the speed of
PAT (or of cytoplasmic streaming) in characeans. These other
mechanochemical motors (dynein–tubulin and kinesin–tubulin) have smaller step lengths (Kimura et al., 2003), as well as
much lower specific reaction rates of the ATPase than do the
myosin–actin systems. Even the fastest specific rate of ATP
hydrolysis, that for the kinesin–tubulin system involved in
anterograde neural transport, is only one-tenth that of the
myosin XI–actin system in Chara (Table 1; Howard, 1997).
If either dynein–tubulin or kinesin–tubulin is to carry auxin
at the observed rate in Chara, there must be an amplification
of the in vitro and in vivo speeds listed in Table 1 (see also
Howard, 1997). Before considering the means by which such
an amplification could occur, we examine the evidence for
the occurrence of tubulin-based mechanochemical motors in
green algae and in embryophytes other than in flagellar motility (dynein) and during mitosis and meiosis (kinesin).
Colchicine, which causes the breakdown of microtubules and thus inhibits, among other processes, dynein- and
kinesin-dependent transport processes, does not inhibit the
myosin–actin-dependent cytoplasmic streaming in Nitella
(Bradley, 1973) at concentrations which are known to disrupt the microtubular apparatus of Nitella (Green, 1962).
Colchicine, but not cytochalasin, inhibited the light-regulated movement of chloroplasts and other cytoplasmic
granules in the coenocytic green alga Dichotomosiphon,
consistent with involvement of dynein or kinesin associated
with microtubules in the movement (Maekawa et al., 1985).
Dichotomosiphon is a member of the class Ulvophyceae, phylogenetically rather distant within the green algae from the
Charophyceae (Graham et al., 2009).
For dynein, contrary to earlier suggestions that flowering
plants had no dynein, it is clear from analysis of genome
sequences that they have genes encoding dyneins despite the
absence of the ‘obvious’ structure which uses dynein, namely
the flagella of male gametes (King, 2002). The Charales have
flagella on their male gametes and so must have at least flagellar dynein, so any mechanism for auxin transport in Chara
based on dynein–tubulin activity is potentially applicable
to embryophytes lacking flagella as well as to the Charales.
Such auxin transport must also, despite its speed, not cause
cytoplasmic streaming since this process relates solely to the
myosin–actin system. The kinesin–tubulin system is involved
in chromosome movement in nuclear division, so the kinesin
family must be represented in all eukaryotes. Presumably it is
the kinesins which bring about axonemal and neural retrograde transport and which have a two orders of magnitude
higher speed of transport (Table 1), which could be available
for carrying auxin as cargo. Some evidence is consistent with
polar movement of auxin involving transport in vesicles for
at least the latter part of the intracellular pathway in Zea
mays L. roots (Schlicht et al., 2006; Mancuso et al., 2007).
However, the only correlation with the cytoskeleton reported
for this pathway is with actin filaments (Schlicht et al., 2006).
If a mechanochemical motor is involved, it is most likely to
be the cytochalasin-sensitive myosin–actin system, which
would not explain the absence of cytochalasin inhibition of
PAT in Chara found by Boot et al. (2012). Vesicle transport
by the Nitella myosin–actin system has been demonstrated by
Kachar et al. (1993; see also Verchot-Lubicz and Goldstein,
2010).
The speed of movement based on a myosin–actin system
can also be increased by having many actin filaments in series,
as in muscle: 100 in series with a speed of 6.3 µm s–1 for each
filament gives a speed of 0.63 mm s–1. However, this mechanism is not appropriate for driving cytoplasmic streaming,
6 | Raven et al.
since it involves contraction depending on overlapping filaments of actin and myosin, followed by relaxation, rather
than continued movement of myosin along actin in the direction of cytoplasmic streaming. This argument can also be
used to rule out having a muscle-like amplification mechanism for dynein–tubulin or kinesin–tubulin.
A further increase in speed of movement at the macroscopic
level is the principle of levers, working at higher levels in more
differentiated organisms. For photosynthetic organisms, the
largest ‘solar mobile’ (an organism whose motility is powered
by photosynthesis: Grassmann, 1988) is the ‘golden jellyfish’
(Mastigias) with symbiotic dinoflagellates in a ‘marine’ lake
in Pulau (Dawson and Hamner, 2003). This organism has a
bell diameter of up to 135 mm, and can swim at up to 70 mm
s–1 by expelling water from the bell by muscular (i.e. myosin–
actin) contraction. Mastigias may not be entirely energized
by photosynthesis: phagomixotrophy is a possible alternative.
The ‘golden jellyfish’ carries out diel vertical migrations (possibly related to obtaining nutrients from the oxycline/thermocline at night and obtaining light energy near the surface
of the turbid upper mixed layer in the day), and horizontal
migrations each day, apparently related to remaining in direct
sunlight during the day in the partly shaded lake (Dawson
and Hamner, 2003).
An analogous mismatch of the speed of movement of a
mechanochemical motor along a cytoskeletal structure is the
dynein–tubulin system of flagella and cilia. Again using a
photosynthetic example, namely the photosynthetic ciliate
Myrionecta rubra Lohman, 1908 (= Mesodinium rubrum),
which is up to 45 µm long and can ‘jump’ at up to 12 mm
s–1, there are contradictory results as to whether cells have
slower, sustained swimming, for example in diel vertical
migration (Lindholm, 1985; Fenchel and Hansen, 2006;
Riisgard and Larsen, 2009). They are photosynthetic, but
rely on eating cryptophyte algae which supply not only their
chloroplasts (kleptoplasty) but also the algal nucleus (karyoklepty) which houses at least 90% of the genes needed to
maintain the plastids. Even so, at least occasional meals are
needed (Johnson, 2011). The most rapid swimmers among
strictly photosynthetic organisms seem to be members of the
freshwater genus Volvox. The spherical colonies of biflagellate green (Chlorophyceae) algal cells are ~1 mm in diameter
and can swim at 1–2 mm s–1. They use this swimming speed
to undergo diel vertical migrations of 10–15 m in stratified
habitats (with vertical water movements of <1–2 mm s–1).
The colonies are nearer the surface in the day when they
can photosynthesize and are deeper at night when they have
access to a higher inorganic nutrient concentration (Beardall
et al., 2009).
The speed ratio (speed of swimming:speed of movement
of dynein along tubulin) for the cilia-based swimming of
M. rubra is >104, while that for flagella-based swimming of
Volvox is ~103. For the muscle-based system in Mastigias,
the speed ratio is ~105. There seem to be no examples of
such amplification of speed for the kinesin–tubulin system.
Thus, in principle, the speed ratio can be explained, although
detailed mechanisms are not all worked out (Howard, 1997;
Lindemann and Lesich, 2010).
Ecological parallels between vertical motile
photosynthetic organisms and cytoplasmic streaming
in the Charales in exploiting habitats with marked
vertical gradients in resource supply
The diel (or longer term) vertical migration enabled by cilia
or flagella, as with those related to changes in the density of
organisms by means of variations in gas vesicle expansion,
the quantity of dense solids, and/or the density of the solution in aqueous vacuoles, has a functional parallel with the
cytoplasmic streaming in characeans with rhizoids in the
dark but high-nutrient sediment, and shoots in low-nutrient
but diurnally illuminated overlying water. In both cases the
motility mechanisms seem to allow the organisms to make
greater use of spatially and temporarily variable resources
(light and nutrients) relative to organisms lacking the motility
mechanisms (Raven, 1981, 1984, 1989, 2003; Beardall et al.,
2009).
Conclusions
The occurrence of PAT in the Charales shows that PAT
originated in the algal ancestors of the embryophytes; further work is needed to determine if PAT occurs elsewhere
in multicellular members of the Charophyceae, in other
multicellular green algae, or even more generally in multicellular algae. In embryophytes, PAT occurs in parenchymatously constructed structures which commonly also have
xylem and phloem (or their bryophyte analogues) as longdistance transport processes in parallel to PAT; in C. corallina PAT shares the pathway for long-distance transport of
nutrients though the parenchymatously constructed nodal
complexes and the single giant cells of the internode and,
in some Chara species, the corticating cells around the
internodal cells. The speed of auxin movement by PAT is
much more rapid than that attributable to diffusion and is
of the same order as the rate of cytoplasmic streaming in
the giant internodal cells, yet complete inhibition of streaming by the inhibitor cytochalasin H does not slow down
auxin transport. Explanations for this independence of PAT
and cytoplasmic streaming are sought in the operation of
other mechanochemical motors, namely dynein–tubulin and
kinesin–tubulin, as alternatives to the myosin–actin system
which powers cytoplasmic streaming. Experiments in which
microtubules are disrupted, for example by colchicine, could
show if one of the tubulin-based motors is involved. If one
of these motors is involved, some mechanism is needed to
amplify the speeds of sliding of the ATPase (dynein or kinesin) to account for the (at least) one order of magnitude
higher speeds seen for auxin transport than for movements
of the ATPase relative to microtubules
Acknowledgements
The author acknowledges the support, and hospitality, of the
University of Technology, Sydney. The University of Dundee
is a registered Scottish charity, No. SC 10596.
Commentary | 7
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