Download Autophagy in Tobacco BY-2 Cells Cultured under

Autophagy in Tobacco BY-2 Cells Cultured under Sucrose
Starvation Conditions: Isolation of the Autolysosome
and its Characterization
Regular Paper
Chihiro Takatsuka1, Yuko Inoue3, Tomoya Higuchi2, Stefan Hillmer4, David G. Robinson4 and
Yuji Moriyasu1,2,*
1
Graduate School of Science and Engineering, Saitama University, Saitama, Japan
School of Science, Saitama University, Saitama, Japan
3
Graduate School of Food and Nutritional Sciences, University of Shizuoka, Shizuoka, Japan
4
Department of Plant Cell Biology, Centre for Organismal Studies, University of Heidelberg, Heidelberg, Germany
*Corresponding author: E-mail, [email protected]; Fax, +81-48-858-3408.
(Received June 3, 2011; Accepted October 7, 2011)
2
Tobacco culture cells carry out a large-scale degradation
of intracellular proteins in order to survive under sucrose
starvation conditions. We have previously suggested that
this bulk degradation of cellular proteins is performed by
autophagy, where autolysosomes formed de novo act as the
major lytic compartments. The digestion process in autolysosomes can be retarded by addition of the cysteine protease
inhibitor E-64c to the culture medium, resulting in the
accumulation of autolysosomes. In the present study,
we have investigated several properties of autolysosomes
in tobacco cells. Electron microscopy showed that the
autolysosomes contain osmiophilic particles, some of
which resemble partially degraded mitochondria. It also
revealed the presence of two kinds of autolysosome precursor structures; one resembled the isolation membrane
and the other the autophagosome of mammalian cells.
Immunofluorescence microscopy showed that autolysosomes contain acid phosphatase, in accordance with cytochemical enzyme analyses by light and electron microscopy
in a previous study. Autolysosomes isolated by cell fractionation on Percoll gradients showed the localization of acid
phosphatase, vacuolar H+-ATPase and cysteine protease.
These results show that starvation-induced autophagy in
tobacco cells follows a macroautophagic-type response
similar to that described for other eukaryotes. However,
our results indicate that, although the plant vacuole is
often described as being equivalent to the lysosome of the
animal cell, a new low pH lytic compartment—the autolysosome—also contributes to proteolytic degradation when
tobacco cells are subjected to sucrose deprivation.
Keywords: Autophagy Autolysosome/autophagosome Cysteine protease Endocytosis Tobacco BY-2 Vacuole.
Abbreviations: Atg, autophagy-related protein; ConcA,
concanamycin A; GFP, green fluorescent protein; 3-MA,
3-methyladenine.
Introduction
Autophagy involves the sequestration of portions of the
cytoplasm including organelles in a membrane-bound compartment followed by the subsequent degradation of the contents in a lytic compartment. This process was first considered
to be an induced response to nutrient depletion, and recent
research on yeast and mammalian cells has put autophagy into
a broader perspective as representing a major degradative
pathway and as contributing to some of the regulation
pathways in cellular homeostasis (Klionsky and Emr 2000).
It is generally accepted that all eukaryotic cells have a lytic
compartment: the lysosome in animal cells, the lysosome/
vacuole in yeast cells and the vacuole in plant cells. By macroautophagy, the cytoplasm to be degraded becomes trapped
within a de novo formed, double-membrane compartment,
the autophagosome. In mammalian cells, the autophagosome
then develops into an autolysosome by acquiring hydrolytic
enzymes and the vacuolar H+-ATPase in the boundary
membrane. This transformation has been considered to be
achieved by vesicle-mediated delivery of newly synthesized
enzymes as well as through fusion to pre-existing primary
lysosomes (Dunn 1994, Eskelinen 2005). In contrast, in yeast,
an autophagosome does not differentiate into a lytic organelle.
Instead, it fuses directly with the vacuole and releases an inner
membrane-bound structure, termed an autophagic body, into
the vacuolar lumen (Klionsky and Ohsumi 1999). The autophagic body is subsequently degraded by vacuolar hydrolases
(Teter et al. 2001).
Microautophagy is a process in which parts of the cytoplasm
are invaginated by the membranes of lysosomes/vacuoles
themselves and then degraded. Such a process has been
observed with the lysosome of mammalian hepatocytes
(Ahlberg and Glaumann 1985) and with the vacuole of the
methylotrophic yeast Pichia pastoris (Tuttle and Dunn 1995,
Sakai et al. 1998).
Plant Cell Physiol. 52(12): 2074–2087 (2011) doi:10.1093/pcp/pcr137, available online at www.pcp.oxfordjournals.org
! The Author 2011. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
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Autophagy in tobacco cells
Autophagic processes that seem to be relevant to developmental processes have been described at the ultrastructural
level in plants on a number of occasions. These include autophagy relating to the biogenesis of lytic vacuoles in vegetative
tissues, protein storage vacuoles in developing seed tissues and
the transformation of protein storage vacuoles to lytic vacuoles
during seed germination (Robinson and Hinz 1997, Herman
and Larkins 1999, Marty 1999, Moriyasu and Hillmer 2000,
Moriyasu and Klionsky 2004, Bassham 2007). However, none
of these cases fits into the classical molds of macro- and microautophagy as given above for mammalian and yeast cells.
On the other hand, it has been thought that suspensioncultured cells perform a form of macroautophagy when
placed under conditions of carbon deprivation (Aubert et al.
1996, Moriyasu and Ohsumi 1996, Rose et al. 2006). However,
the actual lytic compartments in this cell system have not been
characterized (Bassham et al. 2006). Sycamore cells start to
form autophagic ‘vacuoles’ approximately 14 h after transfer
into sucrose-free medium. The majority of these structures
were surrounded by two membranes, and some singlemembrane vesicles with similar contents were observed
inside the vacuole (Aubert et al. 1996). A small number of
autophagic ‘vacuoles’ were bounded by a single membrane
with partially degraded contents. By inhibiting cysteine
protease activity, which increases dramatically upon sucrose
deprivation, the autophagic process is slowed down in tobacco
BY-2 cells, and, as a result, numerous autophagic ‘vacuoles’
1–6 mm in diameter accumulated in the perinuclear cytoplasm
(Moriyasu and Ohsumi 1996). Because the cytoplasmic debris
was still recognizable in these autophagic ‘vacuoles’ they were
termed autolysosomes by the authors. The accumulation and
thus the formation of these autolysosomes were blocked by the
autophagy inhibitor 3-methyladenine (3-MA) (Takatsuka et al.
2004). In Arabidopsis cells cultured in a sucrose-free medium,
the formation of three types of autophagic ‘vacuoles’, which
correspond to autophagosomes to autolysosomes, has been
described (Rose et al. 2006). Taken together, these studies
have shown that macroautophagy accompanying the formation of autophagosomes and/or autolysosomes occurs in
plant cells cultured in nutrient-starved media.
In yeast (Saccharomyces cerevisiae, P. pastoris and Hansenula
polymorpha), 35 autophagy-related proteins (Atgs) have been
identified (Klionsky et al. 2003). Seventeen of these proteins,
which are evolutionarily conserved, play an essential role in
autophagosome formation (Hanaoka et al. 2002, Inoue and
Klionsky 2010). One of these proteins, the ubiquitin-like protein
Atg8, localizes to the membranes of autophagosomes and is
involved in the elongation of autophagosomal membranes in
yeast cells. The formation of autophagosomes can therefore
be monitored using Atg8. Thus, in Arabidopsis and tobacco
cells, autophagosomes have been visualized by the expression
of Atg8 fused to fluoroproteins such as green fluorescent protein (GFP) and yellow fluorescent protein (YFP) (Yoshimoto
et al. 2004, Contento et al. 2005, Thompson et al. 2005,
Toyooka et al. 2006). When transgenic plants were treated
with concanamycin A (ConcA), the structures that derive
from autophagosomes, which appear to correspond to the
autophagic bodies in yeast cells, are seen in the central vacuole
(Yoshimoto et al. 2004, Thompson et al. 2005, Xiong et al.
2007). This result has been taken as evidence to show that
autophagosomes fuse directly with the central vacuole as
occurs in yeast cells (Kirisako et al. 1999). Furthermore, it has
recently been reported that ribulose bisphosphate-containing
bodies, which are formed by the evagination of chloroplasts
into the cytoplasm during senescence, are transported to
the central vacuoles by macroautophagy (Ishida et al. 2008).
In this paper, we establish that starvation-induced macroautophagy in plant cells is qualitatively different from that in
other eukaryotic cells. We show that while autophagosomes
originate as double-membrane-bound structures, the digestion
of ingested cytoplasmic contents occurs to a great extent
before they fuse with the central vacuole. In agreement with
this are the observations that vacuolar H+-ATPase is detectable
in the membrane of the autolysosome, and that hydrolytic
enzymes are present in its lumen.
Results
The characterization of autolysosomes by light
and electron microscopy
Tobacco cells accumulate autolysosomes in a sucrose-free
culture medium containing a cysteine protease inhibitor
(Moriyasu and Ohsumi, 1996). The accumulation of autolysosomes was recognized in almost all cells cultured in sucrose-free
medium containing E-64c using a light microscope equipped
with Nomarski optics (Fig. 1A, arrow), whereas such an accumulation was not seen in cells cultured without the inhibitor
(Fig. 1B). In sections stained with toluidine blue, we found
small (>6 mm in diameter) vacuoles containing densely stained
particles in cells treated with E-64c (Fig. 1C, arrow). Particles
stained with toluidine blue were also seen in cells cultured
without E-64c, but the frequency was lower (Fig. 1D).
Thus autolysosomes in tobacco cells had the appearance
of small vacuoles containing toluidine blue-stained particles.
Electron microscopy revealed that an autolysosome
has a few electron-dense particles (Fig. 1E, arrow). These
electron-dense particles seemed to be the degradation intermediates of parts of the cytoplasm that had been enclosed in
autolysosomes. Indeed, some of the particles were occasionally
less degraded and partially retained the morphology of
double-membrane-bound organelles such as mitochondria or
plastids (Fig. 1F, arrowhead; Supplementary Fig. S1). Fusion
between an autolysosome and the central vacuole was often
observed, suggesting that some autolysosomes fuse with the
vacuolar membrane to release electron-dense contents into
the central vacuole (Fig. 1E). Similar electron-dense particles
were occasionally observed in the central vacuole (Fig. 1E,
arrowhead). Toyooka et al. (2006) also analyzed autophagosomes labeled with a YFP–Atg8 fusion protein in tobacco
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Fig. 1 Light and electron micrographs of tobacco cells cultured under sucrose starvation conditions in the presence of the cysteine protease
inhibitor E-64c. Tobacco cells were cultured in a sucrose-free culture medium for 24 h (A–G) or 10 h (H) in the presence (A, C, E, F, G and H) or
absence (B and D) of the cysteine protease inhibitor E-64c. (A and B) Living cells were observed under a light microscope with Nomarski optics.
(C–H) Cells were fixed, embedded in Spurr’s resin, sectioned and observed by light microscopy (C and D) or by electron microscopy (E–H).
Arrows indicate autolysosomes in A, C and E. Arrowheads indicate partially degraded cytoplasm in the central vacuole in E, partially degraded
organelles such as mitochondria in an autolysosome in F, a putative isolation membrane in G and a putative autophagosome in H. n, nucleus; v,
the central vacuole; m, mitochondrion. Bar: 20 mm in A and B; 10 mm in C and D; 2 mm in E; and 500 nm in F–H.
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Autophagy in tobacco cells
cells under nitrogen starvation by time-lapse video microscopy,
and observed two distinct pathways in which autophagosomes
are delivered to the central vacuole. One is a direct delivery of
autophagosomes to the central vacuole, and the other involves
the engulfment of autophagosomes into small vacuolar structures and subsequent fusion of small vacuoles with the central
vacuole. The small vacuolar structures seem to be autolysosomes, and the latter pathway may correspond to the pathway
we have observed using E-64c in the present study. However,
it should be noted that we observed almost all the particles in
autolysosomes and only a few in the central vacuole, suggesting
that the latter pathway is operating dominantly and that degradation is already well underway before the autolysosomes
fuse with the central vacuole.
In cells cultured under sucrose starvation conditions
without E-64c, we occasionally found a structure resembling
an autolysosome (a small vacuole containing a few electrondense particles) (Supplementary Fig. S2). This suggests that
autolysosomes, which have been recognized in the presence
of E-64c, are real intermediate structures emerging during the
autophagic process in tobacco cells.
In addition to autolysosomes, we found two types of structures that are also probably involved in autophagy (Fig. 1G, H;
arrowhead). One resembles the isolation membrane and the
other the autophagosome of mammalian cells. Both structures
were distinguishable from autolysosomes in that two or more
membranes were present at their boundary and the cytoplasm
enclosed showed no signs of degradation. Similar structures
have also been observed by Toyooka et al. (2006). These
morphological observations suggest that the autophagy that
occurs in tobacco cells cultured under sucrose starvation
proceeds in a manner more similar to mammalian than yeast
macroautophagy.
Immunofluorescence study of autolysosomes
In a previous study, we showed the localization of acid
phosphatase, a marker enzyme of mammalian lysosomes, in
autolysosomes of tobacco cells by enzyme cytochemistry
(Moriyasu and Ohsumi 1996). In the present study, we performed immunofluorescence microscopy on protoplasts
that had accumulated autolysosomes, using an affinity-purified
antibody against acid phosphatase from potato (Fig. 2).
The autolysosomes stained positively, whereas control cells
not accumulating autolysosomes did not show such clear staining (Fig. 2A vs. B). This result provides confirmatory evidence
that autolysosomes contain acid phosphatase. Curiously,
although the central vacuole contains acid phosphatase, it
remained unstained. We assume that this is because the concentration of proteins including acid phosphatase in the central
vacuole is too low to be recognized by immunofluorescence.
Isolation of autolysosomes
Using acid phosphatase as a marker, we attempted to isolate
the autolysosomes of tobacco cells. The cell walls were first
digested to make protoplasts, which were then gently ruptured
and fractionated by isopycnic centrifugation through a Percoll
gradient. All fractions were then analyzed for acid phosphatase
activity. Fig. 3A shows the profile of the activity from control
cells, which were 4-day-old cells immediately after transfer to a
sucrose-free medium. The acid phosphatase activity collected
exclusively in a sharp peak around fraction No. 21. A similar
profile was obtained in the fractionation of the cells that had
been put under sucrose starvation conditions for 24 h without
E-64c (Fig. 3C, filled circle), suggesting that starvation per se
does not alter the intracellular localization of acid phosphatase.
In contrast, the curve of acid phosphatase activity altered very
significantly in the centrifugation tube prepared from the 24 h
starved cells treated with E-64c; another peak (Fig. 3B, arrow)
showed up close to fraction No. 5 near the bottom, in addition
to a peak at around fraction No. 21 (Fig. 3B, filled circle).
This suggested that this peak contained autolysosomes. Thus,
the density of autolysosomes was estimated to be 1.08–1.11 mg
ml–1 using density marker beads (Pharmacia Biotech). Cyt c
oxidase, a marker for mitochondria, and catalase, a marker for
microbodies, were distributed differently from each other.
However, a portion of Cyt c oxidase and almost all of the
catalase activity co-equilibrated in the same peak fractions as
acid phosphatase (Fig. 3B, C). We interpret these data as showing that partially disrupted products of the three organelles,
the central vacuoles, mitochondria and microbodies, collected
in these fractions. The distributions of Cyt c oxidase and
catalase did not change significantly when the cells with
an accumulation of autolysosomes were fractionated
(Fig. 3B vs. C).
Next we analyzed polypeptides in the fractions (Fig. 4).
Acid phosphatase activity in each fraction was measured
again in order to localize the fractions containing autolysosomes (Fig. 4A, arrow). Silver staining of SDS gels showed
that the fractions of autolysosomes (fraction Nos. 4–8) had
many obscure polypeptide bands on a relatively high background (Fig. 4B, left). In contrast, such staining was not seen
in the corresponding fractions from the control cells (Fig. 4B,
right). Thus, some of these polypeptides may represent proteins
of cytoplasmic origin that were enclosed and partially degraded
in autolysosomes. Western blot analysis showed that the fractions of autolysosomes contained acid phosphatase (Fig. 4C),
which is consistent with the results obtained by immunostaining (Fig. 2) and enzyme cytochemistry (Moriyasu and Ohsumi
1996). The autolysosomal fractions also contained the A and B
subunits of vacuolar H+-ATPase (Fig. 4C). This suggests the
presence of a functional H+-ATPase on the membanes
of autolysosomes, which is consistent with their having
an acidic lumen (Moriyasu and Ohsumi 1996). We cannot
exclude, however, the possibility that they originated from
the cytoplasm enclosed in autolysosomes. We also tried to
examine the existence of H+-pyrophosphatase in the autolysosomal fractions, but did not obtain a conclusive result
because H+-pyrophosphatase was detected throughout the
gradient.
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Fig. 2 Immunostaining of tobacco protoplasts for acid phosphatase. Protoplasts were prepared from cells cultured under sucrose starvation for
1 d in the presence (A and C) or absence (B and D) of E-64c. Protoplasts were chemically fixed and bound with anti-potato acid phosphatase
antibody and rhodamine-conjugated secondary antibody. (A and B) Fluorescence images. (C and D) Nomarski images of A and B, respectively.
Significant staining was not observed when the first antibody was omitted. Arrows, autolysosomes. n, nucleus. Bar, 10 mm.
Autolysosomes have cysteine protease activity
E-64c is an inhibitor that binds covalently to papain-type cysteine proteases. Thus cysteine protease activity is not expected
to be recovered in fractions prepared from cells treated with
E-64c. However, leupeptin, which is also able to cause autolysosomes to accumulate (Moriyasu and Ohsumi 1996), inhibits
cysteine protease non-covalently. Thus we expected that if
we isolated autolysosomes from cells treated with leupeptin,
we would be able to remove the inhibitor and detect protease
activity. First we examined whether or not we really could
restore protease activity in the homogenate prepared from
cells treated with leupeptin (Fig. 5). In order to detect protease
activity, we performed activity staining of protease in native
gels containing a standard substrate, gelatin. In the gels, one
main band was detected (Fig. 5, arrowhead). We have previously reported that the protease activity that can be measured
using casein as a substrate increased in response to sucrose
starvation (Moriyasu and Ohsumi 1996). Consistent with this
result, gelatin-degrading activity increased in response to
sucrose starvation (Fig. 5; 0 d vs. 1 d, MeOH and Water).
When the homogenate was prepared from cells that had
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been incubated with E-64c, no protease activity was detected
(Fig. 5; 1 d, E-64c). In contrast, two or more activity bands were
detected in the homogenate prepared from the cells treated
with leupeptin, although the band pattern was different from
the controls, presumably due to the interaction of protease and
leupeptin and their dissociation during electrophoresis (Fig. 5;
1 d, Leu; marked with stars).
To confirm the localization of proteases in isolated autolysosomes, we treated the cells with leupeptin instead of E-64c.
From the profile of acid phosphatase activity, the fractions
of autolysosomes at around fraction No. 6 were identified
(Fig. 6A; left, arrow). In these fractions, we were able to
detect protease activity, consisting of two or more bands on
a native gel (Fig. 6B; left, marked with stars). These bands were
barely detectable when the gels were treated with E-64c after
electrophoresis (data not shown), suggesting that these activities are due to cysteine protease. In contrast, neither protease
activity nor acid phosphatase activity was seen in the corresponding fractions from control cells (Fig. 6A, B; right).
These results support the notion that cysteine proteases are
located in autolysosomes and degrade the enclosed cytoplasm,
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Autophagy in tobacco cells
and that the inhibition of the proteases by E-64c or leupeptin
leads to the accumulation of the degradation intermediates
within autolysosomes.
The autophagy inhibitor 3-MA releases the E-64c
inhibition of endocytosis.
Fig. 3 Fractionation of tobacco cells cultured under sucrose starvation conditions for 1 d in the presence or absence of the cysteine
protease inhibitor E-64c. Tobacco cells were homogenized and
fractionated by density equilibrium centrifugation on a Percoll
gradient. Acid phosphatase (filled circle), catalase (open circle) and
Cyt c oxidase (triangle) activities in each fraction were measured.
Fraction numbers are in the order from the bottom to the top of
centrifuge tubes. (A) Immediately after starvation treatment. (B) After
1 d of culture in the presence of E-64c. The arrow indicates the peak of
the autolysosome fractions. (C) After 1 d of culture in the absence of
E-64c.
E-64c blocks the endocytotic pathway from the plasma membrane to the membrane of the central vacuole (Yano et al.
2004). To confirm this result, we stained the plasma membrane
by incubating the cells with FM4-64 at 0 C and, after washing
out unbound dye, the cells were placed in a fresh culture
medium without the dye at 26 C. Immediately after the start
of culture, the plasma membrane was seen to be heavily
stained, together with a punctate staining in the cell cortex
(Fig. 7A). These punctate structures (Fig. 7A, arrowhead)
were assumed to be endosomes. FM4-64 fluorescence thereafter moved from the plasma membrane to the central vacuole
in cells cultured in a sucrose-free medium for 24 h (Fig. 7B).
In contrast, the majority of the fluorescence associated with
autolysosomes rather than the vacuolar membrane when the
cells were starved for 24 h in the presence of E-64c (Fig. 7C,
arrowhead; see also Fig. 1A in Yano et al. 2004). This result
suggests that some of the membrane for the formation of
autolysosomes flows from the endocytotic pathway leading
to the central vacuole.
3-MA is an inhibitor of autophagy (Gordon and Seglen 1982,
Seglen and Gordon 1982, Takatsuka et al. 2004), but does not
perturb the movement of FM4-64 from the plasma membrane
to the vacuolar membrane (Fig. 7D). We next treated the cells
simultaneously with E-64c and 3-MA. In this situation, autolysosomes did not accumulate (Takatsuka et al. 2004) and
FM4-64 moved normally from the plasma membrane to the
vacuolar membrane (Fig. 7E). Thus, the inhibition of autophagy
by 3-MA abolishes the inhibition of FM4-64 endocytosis by
E-64c.
Yamada et al. (2005) have claimed that the structures accumulating in the presence of E-64d (an esterified form of E-64c)
are endosomes, and proposed that cysteine protease inhibitors
such as E-64c or E-64d inhibit the proteases that are involved in
the fusion of endosomes with the central vacuole, causing the
accumulation of endosomes, which are called autolysosomes in
the present study. It is therefore possible that 3-MA treatment
simply reverses the E-64c inhibition of protease activity, and
thus prevents the accumulation of ‘autolysosomes/endosomes’.
However, measurements of cellular protease activities do not
support this (Fig. 8). First, protease activity measured in cell
homogenates at an acidic pH using casein as a substrate
increased >2-fold in 24 h (Fig. 8A). This was reduced to
approximately 20% by the addition of E-64c to the culture
medium. Inclusion of 3-MA did not perturb the inhibition of
the protease activity by E-64c. Secondly, as shown in Fig. 5, one
major band depicting gelatin-degrading activity in native gels
increased in response to sucrose starvation (Fig. 8B). This
band was not visible in gels prepared from starved cells
which were exposed to E-64c, and the same was observed
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Fig. 4 Western blot analysis of vacuolar proteins in each fraction from tobacco cells cultured under different conditions. Tobacco cells were
cultured under sucrose starvation conditions with (left) or without (right) E-64c for 1 d and were then fractionated on a Percoll gradient. (A) Acid
phosphatase activity in each fraction. Fraction numbers are in the order from the bottom to the top of centrifuge tubes. The arrow indicates the
peak of the autolysosome fractions. (B) Proteins in each fraction were separated by SDS–PAGE and stained with silver. The lane numbers
correspond to the fraction numbers in A. (C) The presence of acid phosphatase (APase), and vacuolar H+-ATPase subunits A (ATPaseA) and B
(ATPaseB) was analyzed by Western blotting. The lane numbers correspond to the fraction numbers in A.
when 3-MA was included. These results show that 3-MA has
nothing to do with the protease levels in cells. Together, the
data strongly suggest that 3-MA releases the inhibition of fusion
between ‘autolysosomes/endosomes’ and the central vacuole
without perturbing the E-64c inhibition of protease activity,
and therefore question the notion that some protease sensitive
to E-64c is required for the fusion between ‘autolysosomes/
endosomes’ and the central vacuole.
Discussion
The exact pathway of macroautophagy is not fully understood
in plant cells. In mammalian cells, an autophagosome fuses with
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a lysosome thereby converting it into an autolysosome.
Similarly in yeast cells, an autophagosome fuses with the vacuole which, since it contains assorted hydrolytic enzymes, is
equivalent to the lysosome. In contrast, in plant cells, it remains
unclear whether an autophagosome fuses directly with the
central vacuole (also a lysosome equivalent) or whether the
autophagosome is transformed into an autolysosome prior to
fusion with the central vacuole. The crucial question is therefore: do autophagosomes become enriched in acid hydrolases?
In previous studies, we showed that special membrane
compartments accumulated in tobacco cells cultured under
sucrose starvation conditions in the presence of a cysteine
protease inhibitor (Moriyasu and Ohsumi 1996, Inoue and
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Autophagy in tobacco cells
Fig. 5 Staining of protease activity in native gels prepared from
tobacco cell homogenates cultured under different conditions.
Tobacco cells cultured under sucrose starvation conditions for 1 d
in the presence of E-64c or leupeptin were homogenized, and proteins
were separated on non-denaturing polyacrylamide gels containing
gelatin as a protease substrate. After electrophoresis, gels were kept
in 50 mM acetate-Na (pH 5.0) containing 2.8 mM 2-mercaptoethanol
at 37 C overnight and then stained with Coomassie brilliant blue.
0 d, immediately after treatment of sucrose starvation; MeOH, cells
cultured for 1 d in the presence of 1% (v/v) methanol as a solvent
control of E-64c addition; Water, cells cultured for 1 d with the
addition of water as a solvent control of leupeptin addition; E-64c,
cells cultured for 1 d in the presence of 10 mM E-64c; Leu, cells cultured
for 1 d in the presence of 10 mM leupeptin. The arrowhead indicates
the main band of protease; the stars mark several activity bands in the
Leu lane.
Moriyasu 2006). These compartments were acidic and
contained acid phosphatase, as was shown by vital staining
with quinacrine and by cytochemical enzyme analysis with
b-glycerophosphate as a substrate. Moreover, the compartments contained electron-dense materials, which assumed
the appearance of partially degraded parts of the cytoplasm.
Interestingly, when mammalian cells executing autophagy are
treated with protease inhibitors such as leupeptin, autolysosomes accumulate in them (Kominami et al. 1983). On the
other hand, when yeast cells under nitrogen starvation conditions are treated with the protease inhibitor phenylmethylsulfonyl fluoride, autophagic bodies, i.e. membrane-bound parts of
the cytoplasm, accumulate in the vacuole (Takeshige et al.
1992). Thus, by analogy to mammalian rather than yeast cells,
it is reasonable to suppose that the compartments accumulated by the inhibition of proteolysis in tobacco cells are autolysosomes. In the present study, we now show by electron
microscopy that the autolysosomes induced in tobacco cells
contained partially degraded organelles such as mitochondria
or plastids (Fig. 1F), confirming that autophagosomes acquire
degradative properties. We have furthermore identified the
precursory structures of the autolysosomes, which appear to
be enclosing a part of the cytoplasm (Fig. 1G) or just have
enclosed it (Fig. 1H). Similar structures have been observed
in sycamore, Arabidopsis and tobacco cells cultured under nutrient starvation conditions (Aubert et al. 1996, Marty 1999,
Rose et al. 2006, Toyooka et al. 2006).
We have successfully isolated cell fractions enriched in autolysosomes and biochemically characterized them. Our results
suggest that autolysosomes have a cysteine protease(s) in their
lumen and most probably an H+-ATPase activity in their membranes. Thus it would seem that autolysosomes in tobacco cells
are able to create and maintain the acidic environment
required for the action of hydrolytic enzymes. This indicates
that autophagosomes acquires hydrolytic enzymes and proton
pump complexes before they coalesce with the central vacuole.
Accordingly, in the autophagy that is performed by tobacco
cells, it is autolysosomes which play a major role in the degradation of the enclosed cytoplasm. A similar conclusion was
reached by Rose et al. (2006) when working on cultured
Arabidopsis cells. The vacuole therefore acts as a recipient
for partially degraded material and completes the degradation.
On the other hand, experiments with ConcA support the
notion that the central vacuole acts as the main lytic compartment. When nitrogen-starved Arabidopsis hypocotyl and root
cells are treated with ConcA, autophagic bodies accumulate
in the lumen of the central vacuole. These are cytoplasmic
droplets (with organelles) surrounded by a single boundary
membrane (Yoshimoto et al. 2004). ConcA is a well-known
inhibitor of vacuolar H+-ATPase and prevents the acidification
of the vacuole interior, implying that autophagic bodies
become visible because they cannot be degraded. However,
in mammalian cells, the effects of ConcA and bafilomycin,
another inhibitor of vacuolar H+-ATPase, seem to be complicated (Klionsky et al. 2008), and in plants it is known that
ConcA also acts at the level of the Golgi apparatus by preventing the formation and release of the trans-Golgi network
(Robinson et al. 2004, Dettmer et al. 2005, Dettmer et al.
2006, Viotti et al., 2010). Thus, although ConcA treatment
does enable an easier detection of autophagy in plant cells, it
obviously distorts the normal autophagic process in plant cells.
How an autophagosome acquires the hydrolytic enzymes
necessary for the degradation of enclosed cytoplasm and
transforms into an autolysosome in tobacco BY-2 cells remains
unclear. It is possible that autophagosomes in tobacco cells are
different from genuine autophagosomes in that they have
hydrolytic enzymes right from the start of their formation.
According to Marty (1978), small vacuoles in root meristematic
cells develop from tubular structures which contain acid phosphatase. These tubules surround a part of the cytoplasm to
form a double-membrane compartment, a kind of autophagosome, which subsequently becomes a vacuole after digestion of
the inner membrane and enclosed cytoplasm.
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Fig. 6 Staining of protease activity in native gels from subcellular fractions obtained from tobacco cells cultured under different conditions.
Tobacco cells were cultured under sucrose starvation conditions for 1 d in the presence (left) or absence (right) of leupeptin. (A) Acid
phosphatase activity in each fraction. The arrow indicates the peak of the autolysosome fractions. (B) Staining of protease activity on native
gels containing gelatin. The lane numbers correspond to the fraction number in A. The stars mark the activity bands of protease.
Another possibility is that autophagosomes acquire hydrolytic enzymes secondarily by an as yet unknown interaction
with the central vacuole. When the vacuolar membranes of
BY-2 cells are labeled with the fluorescent dye FM4-64, the
dye moves to autolysosomes upon shifting the cells to starvation conditions in the presence of E-64c (Yano et al. 2004).
Conversely, the staining intensity of FM4-64 on the vacuolar
membranes decreases. Similarly, a lumenal GFP-tagged vacuolar
protein also moves to autolysosomes, when autolysosomes
are formed (Yano et al. 2004). These results suggest that
there is a contribution of the central vacuole to the formation
of autolysosomes, and explain how they acquire hydrolytic
enzymes from the central vacuole. Interestingly, in the absence
of E-64c, FM4-64 remains on the vacuolar membrane
during starvation-induced autophagy (Yano et al. 2004).
Taken together, it would seem that parts of the vacuolar membrane and lumen join in the formation of autolysosomes,
and autolysosomes once formed return to the central vacuole
in a normal autophagic pathway in the absence of E-64c.
It would appear that the autophagic pathway merges with
the endocytotic pathway in tobacco cells as well as in mammalian cells, since blockage of the autophagic pathway by E-64c
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leads to blockage of the endocytotic pathway (Liou et al. 1997,
Yano et al. 2004). However, it has been proposed that
endosomes accumulate in the presence of cysteine protease
inhibitors because they inhibit the fusion of endosomes with
the central vacuole (Yamada et al. 2005). In contradiction to
this, we now show that the autophagy inhibitor 3-MA overrules
the E-64c block and allows the normal endocytotic pathway to
recover. Thus, the apparent inhibition of endocytosis by E-64c is
due to the inhibition of the autophagic pathway by this
inhibitor. We think that the accumulation of undegraded
materials in autolysosomes makes autolysosomes larger and
hinders their fusion with the central vacuole.
Whether or not the same autophagic pathway as described
here for tobacco BY-2 cells occurs in other plant cells is not yet
known. We have observed young and mature root cells treated
with E-64 d, a more membrane-permeable analog of E-64c, in
barley and Arabidopsis (Moriyasu et al. 2003, Inoue et al. 2006).
In these cells, partially degraded materials derived from the
cytoplasm appeared to accumulate in large vacuoles and
small vacuoles such as autolysosomes. The detailed observation
of autolysosomes and the central vacuole is now in progress in
our laboratory.
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Autophagy in tobacco cells
Materials and Methods
Plant material
Tobacco culture cells (BY-2) were cultured in a 300 ml
Ehrenmeyer flask at 26 ± 1 C with rotation of 110 r.p.m. The
culture was maintained by transferring 1.5 ml of cell suspension
at the stationary phase to 80 ml of fresh culture medium once
a week. The culture medium consisted of water, 4.3 g l–1
Murashige and Skoog salt mixture (M7150, Sigma), 3% (w/v)
sucrose, 0.2 mg l–1 2,4-D, 2 mg l–1 glycine, 100 mg l–1
myo-inositol, 0.5 mg l–1 nicotinic acid, 0.5 mg l–1 pyridoxineHCl and 0.1 mg l–1 thiamine-HCl. A sucrose-free culture
medium was prepared by omitting sucrose from the culture
medium and was used for putting cells under carbon starvation
conditions. The pH of these culture media was adjusted
to between 5.8 and 6.0 with fresh KOH.
Sucrose starvation
Four-day-old cells, which were at the logarithmic phase, were
collected by centrifugation at 500g for 5 min. The cells were
suspended in the original volume of the sucrose-free culture
medium and precipitated again by centrifugation. They were
suspended again in the same volume of fresh sucrose-free
culture medium and kept at 26 ± 1 C with rotation of
110 r.p.m.
When the cells were treated with E-64c (a gift from Taisho
Pharmaceutical Company), one-hundredth of the culture
medium of 1 mM E-64c dissolved in methanol was added to
the cell culture except in experiments for cell fractionation.
Thus the final concentration of E-64c was 10 mM. As a control
of solvent effects, the same volume of methanol alone
was added. Thus the final concentration of methanol was
1% (v/v).
Microscopy
In order to observe whole living cells and immunostained
protoplasts, a light microscope (OptiPhoto, Nikon) that is
fitted with Nomarski interference optics and an epifluorescence
apparatus was used. Photographs were taken through a 20
or 40 objective lens and a 5 projection lens using color film
(Fujicolor ASA400, Fuji). To observe cells stained with FM4-64,
confocal laser microscopes (MRC-1024, Bio-Rad, or FV1000-D,
Olympus) were used.
For sectioning, the cells were embedded in Spurr’s resin.
They were fixed with 2% (w/v) glutaraldehyde and 1% (w/v)
formaldehyde in 0.1 M sodium cacodylate-HCl (pH 6.9) at room
temperature for 1 h and at 4 C overnight, and post-fixed with
Fig. 7 The effect of the autophagy inhibitor 3-MA on the endocytosis
of FM4-64. Plasma membranes of tobacco cells were stained with
FM4-64, and endocytosis was observed. (A) Immediately after staining
with FM4-64. (B) After 1 d in the absence of E-64c and 3-MA. (C) After
1 d in the presence of E-64c alone. (D) After 1 d in the presence of
3-MA alone. (E) After 1 d in the presence of both E-64c and 3-MA.
Arrowheads indicate putative endosomes in A and the accumulation
of autolysosomes/endosomes in C. n, nucleus. Bar, 50 mm.
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Immunofluorescence staining for acid
phosphatase
Fig. 8 The effect of the autophagy inhibitor 3-MA on intracellular
protease activities in tobacco cells. After tobacco cells were cultured
in a sucrose-free medium for 1 d in the presence or absence of E-64c
and 3-MA, the cells were homogenized. (A) Protease activity in the
homogenate was measured using casein as a substrate. Values relative
to the value at 0 d are shown. 0 d, immediately after culture in a
sucrose-free medium; 1 d, 1 d after culture in the absence of E-64c and
3-MA; 1 d+E64, 1 d after culture in the presence of E-64c alone;
1 d+3MA+E64, 1 d after culture in the presence of both 3-MA and
E-64c. (B) Activity staining of protease on a native polyacrylamide
gel. + and – mean the presence and absence of the respective
inhibitors.
1% (w/v) osmium tetroxide at room temperature for 2 h.
After they were immersed in 1% (w/v) uranyl acetate at
room temperature for 2 h, the cells were dehydrated with
a series of ethanol/water mixtures and propylene oxide. They
were then embedded in the resin. Sections of 1 mm thick
were made from the blocks and stained with toluidine
blue. They were observed by light microsocpy (OptiPhoto,
Nikon). Sections of approximately 70 nm thick were made
from the blocks and stained with uranyl acetate and lead nitrate. They were observed by electron microscopy (H-7000,
Hitachi).
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To make protoplasts, cells were collected by centrifugation
and kept at 28 C for 60 min in 10 mM MES-Na buffer
(pH 5.5) containing 1% (w/v) Cellulase Onozuka RS (Yakult
Pharmaceutical Industry Co., Ltd.), 0.1% (w/v) Pectolyase Y-23
(Seishin Pharmaceutical Co., Ltd.) and 0.4 M sorbitol, and protoplasts were collected by centrifugation at 500g for 5 min,
washed with 0.4 M sorbitol, and fixed at room temperature
for 1 h with 50 mM phosphate-Na (pH 7.0) containing 3.7%
(w/v) formaldehyde, 5 mM EGTA, 3% (w/v) sorbitol and
0.02% (w/v) sodium azide. After the fixative was exchanged
with a fresh one, protoplasts were stored at 4 C until the
next step. In the next step, the protoplasts were washed with
50 mM phosphate-Na buffer (pH 7.0) containing 5 mM EGTA,
3% (w/v) sorbitol and 0.02% (w/v) NaN3 for 5 min three times
and for 1 h once. They were then treated with 0.5% (w/v)
Triton X-100 for 5 min at room temperature. After they were
washed with the same buffer for 5 min three times, the
protoplasts were kept for 30 min in blocking solution
{phosphate-buffered saline [P-3813, Sigma; 10 mM phosphate
buffer (pH 7.4), 138 mM NaCl, 2.7 mM KCl], 0.25% (w/v) bovine
serum albumin (BSA), 0.25% (w/v) gelatin and 0.05% (w/v)
NP-40}. The blocking solution was removed and the protoplasts
were incubated overnight at 4 C in the blocking solution
containing antibody against acid phosphatase. The protoplasts
were washed with the blocking solution for 5 min three times
and then incubated with the blocking solution containing
1/400 diluted anti-rabbit IgG–tetramethylrhodamine isothiocyanate conjugate (T-6778, Sigma) for 1 h at room temperature.
After the protoplasts were washed with the blocking solution
three times each for 5 min, they were observed by epifluorescence microscopy.
Antibody against acid phosphatase used in this staining was
affinity-purified using nitrocellulose paper blotted with acid
phosphatase from potato according to the method by Pringle
et al. (1991). Briefly, acid phosphatase from potato (P-1146,
Sigma) was separated by SDS–PAGE and transferred electrophoretically onto nitrocellulose paper. The membrane strip
with the major band of potato acid phosphatase (approximately 45 kDa) was cut out and used as an affinity purification
matrix.
Cell fractionation
Four-day-old cells in 80 ml of culture medium were precipitated
by centrifugation at 500g for 5 min. Supernatant was removed
and the cells were suspended in 80 ml of sucrose-free culture
medium. Cells were precipitated again and the supernatant was
removed. Cells were then suspended in 80 mk of fresh
sucrose-free culture medium. After 80 ml of 10 mM E-64c or
methanol (as a solvent control) was added, the suspensions
were cultured at 26 C with rotation of 110 r.p.m. Immediately
or 24 h after sucrose starvation, cells were collected by centrifugation at 500g for 5 min. To prepare protoplasts, cells were
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Autophagy in tobacco cells
washed once with 0.4 M sorbitol and kept in 80 ml of 10 mM
MES-Na buffer (pH 5.5) containing 1% (w/v) Cellulase Onozuka
RS, 0.1% (w/v) Pectolyase Y-23 and 0.4 M sorbitol at 26 C for 1 h
with rotation. The resulting protoplasts were collected by
centrifugation at 500g for 10 min, and the supernatant was
discarded. Protoplasts were suspended in 0.4 M sorbitol and
pelleted by centrifugation again. They were then disrupted in
10 ml of 0.4 M sorbitol containing 50 mM HEPES-K (pH 7.5),
1 mM EDTA, 10 mM leupeptin (Peptide Institute), 10 mM
pepstatin A (Peptide Institute) and 1 mM 4-(2-aminoethyl)
benzenesulfonyl fluoride hydrochloride by passing through a
needle. Organelles that were released from the cells were
pelleted by centrifugation at 17,000g (JA-20 rotar, Beckman)
for 5 min. The pellet was suspended in 5 ml of 0.4 M sorbitol
containing 10 mM HEPES-K (pH 7.5) and 1 mM EDTA and then
filtered through nylon mesh (mesh size: 10 mm). The filtrate
was put on 60 ml of 0.4 M sorbitol containing 30% (v/v)
Percoll (Amerciam-Pharmacia), 10 mM HEPES-K (pH 7.5)
and 1 mM EDTA, and then centrifuged at 50,000g for 1 h
(RP45T rotar, Hitachi) to carry out density gradient and organelle separation simultaneously. After centrifugation, the content of a centrifuge tube was divided into about 30 fractions
(each 2 ml).
Acid phosphatase activity in each fraction was measured in a
mixture of 500 ml of 0.1 M acetate-Na (pH 5.0), 380 ml of water,
100 ml of each fraction and 20 ml of 50 mM p-nitrophosphate.
The reaction was performed at 37 C for 20 min, and then
stopped by the addition of 1.6 ml of 1 M Na2CO3. After
Percoll in the reaction mixture was precipitated by centrifugation at 1,000 r.p.m. for 5 min, the A405 of the supernatant
was measured.
To measure Cyt c oxidase activity, a marker of mitochondria,
in each fraction, 100 ml of each fraction was mixed with 100 ml of
4 mg ml–1 digitonin. The mixture (50 ml) was mixed with 900 ml
of Cyt c substrate solution consisting of 26.4 mg of Cyt c and
28 mg of Na2S2O3 dissolved in 40 ml of 100 mM phosphate-K
(pH 7.0) in a cuvette, and then the decrease in the A240 at 37 C
was monitored.
To measure catalase activity, a marker of microbodies, in
each fraction, 1 ml of 10 mM H2O2 in 50 mM phosphate-K
(pH 7.0) was mixed with 50 ml of each fraction in a
cuvette, and then the decrease in the A240 was monitored
at 37 C.
(w/v) skim milk, 150 mM NaCl and 0.05% (w/v) Tween-20
at 4 C overnight.
Anti-acid phosphatase (potato) antibody (10 mg ml–1,
Polyscience) was used at 1 : 3,000 dilution without further purification; anti-A and B subunits of vacuolar H+-ATPase antibody
and anti-H+-pyrophosphatase antibody (generous gifts from
Dr. Maeshima, Nagoya University) were at 1 : 2,500 dilution.
The secondary antibody used was anti-rabbit IgG antibody
conjugated with alkaline phosphatase (S3731, Promega). The
binding of the primary and secondary antibodies to membranes
was done in 10 mM Tris–HCl (pH 8.0) containing 150 mM NaCl
and 0.05% Tween-20. Nitroblue tetrazolium and 5-bromo-4chloro-3-indolyl phosphate were used to detect the activity.
Activity staining of gelatin-degrading activity
on a polyacrylamide gel
A polyacrylamide gel [7.5% (w/v)] containing 0.04% (w/v)
gelatin was made by polymerizing the mixture of acrylamide
and methylene-bis-acrylamide together with gelatin. Cell
homogenate, prepared in the same way as for measuring
casein-degrading activity, was mixed with one-fifth volume of
20% (w/v) sucrose and 0.01% (w/v) bromophenol blue and run
on the gel at 4 C according to Davis (1964). After electrophoresis, the gel was washed three times with 20 mM MES-Na
(pH 5.5) containing 2.8 mM 2-mercaptoethanol, and incubated
in the same buffer at 37 C overnight. The gel was thereafter
stained with Coomassie Brilliant Blue, destained and
photographed.
Endocytosis of FM4-64
Four-day-old culture (5 ml) was centrifuged at 500 r.p.m. for
5 min and the sedimented cells were suspended in 5 ml of
sucrose-free culture medium. The suspension was centrifuged
again and the sedimented cells were resuspended in 5 ml of
sucrose-free culture medium. To 1 ml of cell suspension in
the sucrose-free culture medium, 10 ml of 1 mM E-64c or
methanol alone and 0.5 ml of 0.1 M 3-MA or water alone
were added. Immediately, 100 ml of the suspension was put
into a polypropylene tube (12 75 mm) and cooled down to
0 C. To this tube, 1 ml of 10 mM FM4-64 (Molecular Probes)
solution in dimethylsulfoxide (DMSO) was added, and the
culture was kept at 0 C for 20 min. FM4-64 was washed
off and the culture was returned to 26 C; this was defined
as t = 0.
SDS–PAGE and western blotting
An aliquot of each fraction was mixed with the same volume of
sample preparation buffer (Tris–SDS–bME, Enprotech), and
then heated at 100 C for 5 min. Proteins were separated on
10% polyacrylamide gels by SDS–PAGE according to Laemmli
(1970). After electrophoresis, gels were stained with silver.
Proteins in another gel were electrotransferred onto a nitrocellulose membrane in a solution consisting of 25 mM Tris,
192 mM glycine and 20% methanol. After transfer, membranes
were immersed in 10 mM Tris–HCl (pH 8.0) containing 5%
Measurement of casein-degrading activity
The cells were homogenized and casein-degrading activity was
measured in almost the same way as reported previously
(Moriyasu and Inoue 2008). Cells in 3 ml of culture were collected by centrifugation at 500g for 5 min. After the cells were
washed with 3 ml of sucrose-free culture medium, they were
homogenized at 4 C with 100 mM MES-Na (pH 5.5) containing
28 mM 2-mercaptoethanol using a mortar-driven Teflon homogenizer. The homogenate was centrifuged at 15,000 r.p.m. for
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C. Takatsuka et al
10 min. The supernatant (60 ml) was mixed with 40 ml of 0.5%
(w/v) fluorescein isothiocyanate-labeled casein solution,
which had been prepared according to Twinning (1984).
Supplementary data
Supplementary data are available at PCP online.
Funding
This work was supported by the Institute for Space
and Aeronautical Sciences [Grant for Basic Biology Study
oriented to utilization of Space station]; Japan Space
Forum [‘Ground-based Research Announcement for Space
Utilization’ to Y.M.]; Japan Society for the Promotion of
Science; the Ministry of Education, Culture, Sports, Science
and Technology [Grants-in-Aid for Scientific Research
23120504 and 23570222 to Y.M.].
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