Free-flow electrophoresis for purification of plant mitochondria by surface charge

TECHNICAL ADVANCE

Free-flow electrophoresis for purification of plantmitochondria by surface charge

Holger Eubel1, Chun Pong Lee1,2, John Kuo3, Etienne H. Meyer1, Nicolas L. Taylor1 and A. Harvey Millar1,*

1ARC Centre of Excellence in Plant Energy Biology, The University of Western Australia, 35 Stirling Highway, M316,

Crawley, WA 6009, Australia,2School of Biomedical, Biomolecular and Chemical Sciences, The University of Western Australia, Stirling Highway,

M310, Crawley, WA 6009, Australia, and3Centre for Microscopy, Characterisation & Analysis, The University of Western Australia, 35 Stirling Highway, M010,

Crawley, WA 6009, Australia

Received 1 February 2007; revised 2 July 2007; accepted 4 July 2007.*For correspondence (fax +61 8 6488 4401; e-mail [email protected]).

Summary

Sample purity is the key for a successful in-depth analysis of any given subcellular proteome. The suitability of

free-flow electrophoresis to assist conventional, centrifugation-based techniques in the preparation of plant

mitochondria from green and non-green tissue was assessed by various means, including functional assays,

immunoblots, electron microscopy and differential gel electrophoresis. Results indicated a significant increase

in purity of the mitochondrial samples, highlighted specific contaminants previously reported as mitochondrial

proteins, and also pointed to new means for separating plastids and peroxisomes from mitochondria in plant

organellar extracts by exploiting differences in surface charge. This approach has the potential to allow a

deeper and more comprehensive investigation of the Arabidopsis organellar proteomes, by providing a second

dimension of separation based on surface charge in addition to conventional centrifugation purification

protocols relying on size and density.

Keywords: plant mitochondria, proteomics, sample purity, organellar contamination, free-flow electrophoresis,

centrifugation.

Introduction

The purification of a subcellular organelle to the highest

possible level is desirable for many experiments. Subcellu-

lar proteomics is particularly dependant on the purity of the

starting material. As a result of advances in mass spec-

trometry, coverage of low-abundance proteins in samples is

increasing and often leads to the discovery of impurities in

organellar fractions. One solution has been to compare less

enriched fractions of a certain compartment with more

enriched ones, and to measure the increase or decrease in

the abundance of a protein to use it as an indicator for its

intracellular localization (Dunkley et al., 2004, 2006; Nelson

et al., 2006). Although this provides an elegant way to ana-

lyze those compartments that cannot be purified to a high

degree for technical reasons, the depth in the analysis

appears to suffer. Therefore, if feasible, the purification of a

cellular compartment to the highest possible degree is pre-

ferred over a differential approach, but the combination of

both is likely to be most beneficial.

Plant mitochondria have been analyzed frequently

by standard proteomic techniques (Bardel et al., 2002;

Heazlewood et al., 2003, 2004; Kruft et al., 2001; Millar et al.,

2001). But as the number of proteins identified so far in these

organelles is significantly lower than expected, from analysis

of the Arabidopsis nuclear genome by targeted prediction

programs (summarized in Heazlewood et al., 2005), many

more proteins probably remain experimentally unidentified.

Additionally, according to the compilation of data in the

SubCellular Proteomic Database for Arabidopsis (SUBA:

http://www.suba.bcs.uwa.edu.au) (Heazlewood et al., 2007),

some proteins assigned to mitochondria by these studies

ª 2007 The Authors 583Journal compilation ª 2007 Blackwell Publishing Ltd

The Plant Journal (2007) 52, 583–594 doi: 10.1111/j.1365-313X.2007.03253.x

are also claimed in other cell organelles by means of mass

spectrometry and GFP analysis. These proteins may repre-

sent organellar contaminations or proteins targeted to more

than one organelle.

Differential and isopycnic centrifugation using sucrose or

Percoll gradients to separate organelles on the basis of size

and density have been traditionally used in isolation of

mitochondria from plant tissues (Day et al., 1985; Leaver

et al., 1983; Neuburger et al., 1982). These approaches have

also been used in the preparation of mitochondria for

proteomic analysis (Bardel et al., 2002; Heazlewood et al.,

2004; Kruft et al., 2001; Millar et al., 2001). Unfortunately,

based on these parameters, biological variation will always

lead to a degree of overlap between different organelles or

compartments. Free-flow electrophoresis in zone electro-

phoresis mode (ZE-FFE) does not rely on size or density, but

provides separation electrophoretically by using differences

in the surface charge of the organelles. ZE-FFE has success-

fully been employed to purify different cell populations of

the immune system (Baier et al., 1988; Hansen and Hannig,

1982; Hansen et al., 1989; Heidrich and Hannig, 1989), as well

as subcellular particles like endosomes, lysosomes, peroxi-

somes and endoplasmic reticulum (ER) vesicles (Marsh,

1989; Morre et al., 1989; Schmid et al., 1988; Volkl et al.,

1997). To our knowledge, ZE-FFE has been reported once on

plant material for the production of highly purified vesicles

of the Arabidopsis plasma membrane and the tonoplast

(Bardy et al., 1998). Recently, ZE-FFE has been used to purify

yeast mitochondria to an increased homogeneity (Zischka

et al., 2003, 2006). The presence of plastids in plants adds

an additional challenge for any mitochondrial purification

technique, as plastids are known to be the major source of

contamination for mitochondria isolated from photosyn-

thetic material. Here, we present a study employing ZE-FFE

for the purification of plant mitochondria from green

and non-green material, and demonstrate the value of

this approach by identifying impurities in mitochondrial

preparations.

Results and discussion

Visual analysis of the organelle separation

process during FFE

The starting material for FFE separations consisted of an

enriched mitochondrial fraction, produced by differential

centrifugation and Percoll density gradient centrifugation,

from either dark-grown Arabidopsis cell culture or light-

grown Arabidopsis shoots from hydroponic cultures.

Washed and Percoll-free organelles were then injected into

the separation chamber of the instrument, which features

a Perspex top cover. This allowed the observation of the

separation process of the light-scattering, translucent

organelles. At the front end of the chamber, in close

proximity to the cathode stabilization medium, the sample

was injected into the separation chamber (Figure 1, lower

panels). It was taken up by a stream of separation medium

and transported towards the rear end of the chamber. As

soon as the sample entered the zone of the electric field it

became deflected towards the anode. The formerly

homogenous sample stream spread out in a fan-shaped

manner. Usually, two main streams were observed, a

green one to the left and a white/yellow one to the right.

Most probably, the green stream represented the plastid

fraction of the sample (1st streams in Figure 1), whereas

the white/yellow stream represented the bulk of the

(a) (b) Figure 1. Separation of pre-purified mitochon-

drial fractions from Arabidopsis cell cultures (a)

and shoot hydroponic cultures (b) by free-flow

electrophoresis (FFE) (lower panels) into 96-well

plates. The absorbance at a wavelength of

280 nm was measured (upper panels). Arrows

and protein peaks 1–4 represent the four most

abundant and reproducible organelle popula-

tions. SI, sample inlet; ), cathode of the separa-

tion chamber; +, anode of the separation

chamber. Arrows at the bottom represent the

seven media inlets to the separation chamber;

arrows on top represent the 96 outlets of the

fraction collection device. The zones of the

electric fields in the lower panels are colored in

grey.

584 Holger Eubel et al.

ª 2007 The AuthorsJournal compilation ª 2007 Blackwell Publishing Ltd, The Plant Journal, (2007), 52, 583–594

material and was formed by mitochondria (2nd streams in

Figure 1). Often minor streams of both colors can be

observed to the left, to the right and in-between the two

major streams. Each of these minor streams is sharply

focused, and we therefore suspect that they consist of

other organelles, defined associations of organelles or

distinct organelle subpopulations with different surface

charges. Analysis of the minor streams is difficult, because

the sharp resolution observed in the separation chamber is

not reflected by the protein-concentration in the wells of

the 96-well plates used to collect the fractions (Figure 1).

Here, broader distributions were observed that were partly

overlapping. Two minor streams are most often observed,

represented by the 3rd streams in Figure 1 running close

to mitochondria, and the 4th streams that were the least

deflected through the chamber and are usually not sharply

focused by FFE.

General distribution of proteins in FFE fraction

As can be seen in Figure 1 (upper panels), the bulk of the

protein (2nd streams) is deflected during electrophoresis

and is collected on the anionic side of the fraction that is

in-line with the sample injection point. Additional peaks

(from the 1st, 3rd and 4th streams) can be observed to the

left and the right of the major peak, spreading over an area of

40–50 fractions. Thus, assuming that the organelles are still

intact after electrophoresis, it can be concluded that the

cellular components of an enriched mitochondrial fraction

are actually separated during the FFE treatment. Because the

sample consists of an enriched mitochondrial fraction, the

major protein peaks (from 2nd streams) are expected to

represent the mitochondrial components of the samples.

The other peaks are thus suspected to be contaminating cell

components.

Some obvious differences exist between the samples

derived from the cell culture (Figure 1a) and shoot material

(Figure 1b). In shoots, fractions on the anodic side of the

major peak have considerably higher protein contents than

those of the cell culture. Although maintained in the light,

bulking up of the cell culture for the mitochondria prepara-

tion took place in the dark to reduce chloroplast contamina-

tion. Therefore, a major difference between the two samples

is likely to be the level of chloroplast material within the

cells. One can speculate that it is these organelles which are

responsible for the differences in protein concentration on

the anodic side of the main peak between the two samples.

The color of the collected FFE fractions supports this

hypothesis: green fractions are found closer to the anode

(representing the areas of the 1st streams) than those found

in the areas of the 2nd streams, which had a yellow/brown

color (data not shown). The size of peak 3 is another minor

difference between the two samples as it is pronounced in

the cell culture derived sample.

Distribution of marker proteins in FFE fractions

To further establish the distribution of organelles across the

FFE fractions, an aliquot of every second fraction was loaded

onto one-dimensional SDS gels to perform western blots.

Antibodies raised against protein markers for mitochondria,

plastids and peroxisomes were used to probe the blots

(Figure 2).

A monoclonal antibody against maize mitochondrial

HSP70 recognized a protein with a molecular weight of

about 75 kDa, which fits very well with the calculated

molecular weight for this protein (73 kDa). For the cell

culture as well as the shoots, mitochondrial HSP70 is present

in only those fractions containing the bulk of the protein

content, supporting the hypothesis that these fractions

comprise the mitochondrial portion of the sample.

Polyclonal antibodies recognizing the small subunit of

RubisCO (RbcS) detected a protein of 15–16 kDa, which is in

line with the observed molecular weight for the mature

Figure 2. One-dimensional SDS–PAGE and im-

munoblots of every second of the central 30

fractions collected after free-flow electrophoresis

(FFE). The gels were loaded on a volume basis to

monitor the distribution of three marker proteins

for mitochondria (mtHSP70), peroxisomes (3-

ketoacyl-CoA thiolase, Kat2) and plastids (small

subunit of RubisCO, RbcS).

Isolation of plant mitochondria by FFE 585


protein (Peltier et al., 2006). Only a weak signal could be

obtained with the antibody for the cell culture. Those

fractions reacting with the antibody ran to the left of the

mitochondrial fractions, closer to the anode. In the shoot

sample, significantly more fractions contained RbcS and in

higher concentrations. Again, these results support the

assumptions made on the basis of the A280-based protein

concentrations of the individual FFE fractions (Figure 1) and

the visual inspection of fraction colors in the FFE run.

In contrast to plastids, peroxisomes seem to have a

slightly lower electrophoretic mobility than mitochondria.

Polyclonal antibodies directed against peroxisomal 3-keto-

acyl-CoA thiolase (Kat2) recognized a band of approximately

47 kDa, matching the calculated molecular weight of

48.5 kDa for this protein. In both samples, peroxisomes

seem to migrate closely to the mitochondria, thereby

creating a certain overlap of fractions containing both

marker proteins. This overlap is more pronounced in the

shoots than in the cell culture, probably resulting from

different physical properties of this organelle in the two

samples. By picking only the fraction with the highest

protein concentration, a reduction in peroxisomal conta-

mination can still be achieved.

Composition of the 4th peak found in FFE separation

Whereas the identity of the three most deflected protein

peaks (1st, 2nd and 3rd streams in Figure 1) has been

established by means of marker proteins, the identity of the

fourth peak could not be unambiguously revealed by these

methods. This fourth stream was not consistent as its

abundance varied between individual FFE runs for unknown

reasons; typically it represented 2–10% of the integrated 280-

nm absorbing material (Figure 1). The protein content of this

peak was separated on a one-dimensional SDS gel (data not

shown) and major bands were analyzed by LC-MS/MS. The

top three hits for each band are listed in Table S1. These are

major proteins localized in or associated with the mito-

chondrial or peroxisomal membranes based on comparison

with mitochondrial proteome studies and targeting predic-

tions (Heazlewood et al., 2004, 2007). The bulk of the evi-

dence thus suggests this peak consists of ruptured organelle

membranes.

Purity and homogeneity of the final isolate

As shown in Figure 2, mitochondria isolated from cell

culture or shoots differ in their main contaminants. In cell

cultures peroxisomes are the major contaminants, whereas

in shoots the chloroplast contamination dominates over

peroxisomal contamination.

To reduce the loss in yield after FFE, fractions with the

highest mitochondria concentration were selected by eye

and pooled. For the shoot mitochondria, this meant a

peroxisomal contamination level that was probably higher

than it would be if pooling was performed after western blot

analysis, but was still lower compared with the level of

contamination before FFE (Figure 3a). For the cell culture,

only the two fractions that represented the anodic side of the

mitochondrial peak were pooled, to minimize peroxisomal

contaminants from the cell culture sample.

The level of purity of the final mitochondrial isolates has

been further assessed in various ways. For the cell culture,

the degree of peroxisomal contamination was ascertained

by catalase activity. As can be seen in Figure 3(b), FFE-

purified mitochondria from cell culture produce oxygen at a

much lower rate. On average, a 5- to 6-fold reduction in

peroxisomal soluble protein contamination of the mitochon-

drial fraction can be achieved through FFE. For shoot

material, a visual indication of a reduction in the degree of

plastidic contamination from fractions pooled after FFE was

the green color of the mitochondrial pellet. In Figure 3(c)

organellar pellets derived from Arabidopsis shoots before

and after FFE are compared. Whereas the pellet before the

FFE run still incorporates green material, it is nearly devoid

of it after the FFE treatment. Chlorophyll assays showed that

this reduction was more than threefold on a protein basis

(Figure 3c). These findings are also reflected in immuno-

blots conducted with the pooled mitochondrial fractions

derived from shoot or cell culture material (Figure 3a). The

largest improvement is found for the shoot sample, where

FFE separation led to a nearly complete loss in the signal for

the plastid marker, RbcS. The signal for the peroxisomal

marker, Kat2, is also clearly reduced after FFE. Similar results

have also been obtained for the cell culture sample.

Although the immunoblot is faint for RbcS, a reduction in

the intensity was observed after FFE. However, the major

contaminants in these samples were peroxisomes, and after

FFE the Kat2 immunoblot intensity was markedly decreased.

A broader approach was also undertaken to investigate

the success of FFE purification more systematically. Differ-

ential two-dimensional iso-electric focusing (IEF)/SDS-PAGE

using the fluorescent dyes Cy2, Cy3 and Cy5 was performed

for three independent cell culture sample sets (Figure 4a)

and shoot samples sets (Figure 4b). Overall, the general

protein spot patterns are very consistent with the mitochon-

drial proteome profiles previously published (Millar et al.,

2001). A threshold of at least a threefold change in protein

abundance was considered to be a significant effect, and

was used to identify proteins that changed in abundance

following FFE treatment. In each case the changing spots

were more abundant in the pre-FFE fraction. Among this set

of proteins, the most abundant spots on Coomassie stained

gels were selected and analyzed using LC-MS/MS (indicated

by arrows in Figure 4). The identities of the proteins are

summarized in Tables 1 and 2. It was apparent that for some

spots the staining intensity was different for fluorophore

labeling and Coomassie staining. Therefore, not all of the



major spots in Figure 4 have been analysed because they

could not be identified with confidence from the Coomassie

stained gel.

For the cell culture samples, 22 spots were analyzed, and

all of them were found to be more abundant in the pre-FFE

sample. The 22 matches were of non-mitochondrial origin

and matched to known proteins from plastids, peroxisomes,

the cytosol or the nucleus (Table 1). Twenty spots from the

shoot mitochondrial samples with at least a threefold

change in abundance were analyzed by LC-MS/MS, resulting

in the identification of 23 different gene products (Table 2).

Two identified proteins (subunit 6 of the succinate dehydro-

genase and the beta subunit of ATP-synthase) are well-

known mitochondrial proteins, but both were secondary hits

and are not likely to be the source of the detected change in

abundance. The bulk of the identified proteins are located in

the plastid, with two in the ER. Therefore, in both samples,

various proteins of non-mitochondrial origin were present

before FFE, but were substantially decreased in abundance

following FFE.

In combination these data clearly indicate a substantial

gain in purity of the mitochondrial fraction by means of FFE.

The identified putative contaminants in mitochondrial

preparations provided by Tables 1 and 2 also give several

(a)

(b) (c)

Figure 3. Assessment of the purity of pooled

mitochondrial fractions.

(a) One-dimensional SDS–PAGE and immuno-

blots of organelles from cell culture and hydro-

ponically grown shoot material before and after

free-flow electrophoresis (FFE). A total of 10 lg

of protein has been loaded per lane. Immuno-

blots are shown for mitochondria (mtHSP70),

peroxisomes (3-ketoacyl-CoA thiolase, Kat2) and

plastids (small subunit of RubisCO, RbcS).

(b) Catalase activity measured in mitochondria-

enriched fractions from cell culture before and

after FFE. The arrow above the graph indicates

the point of H2O2 injection. Rates are given in

nmol O2 min)1 mg)1 protein (mean � SD,

n = 3).

(c) Photographs showing the appearance and

colour of mitochondrial pellets derived from

hydroponically grown shoot material before

and after FFE. The results of replicated chloro-

phyll assays (ng mg)1 protein) are given below

(mean � SD, n = 3).



(a)

(b)

Figure 4. DIGE two-dimensional iso-electric

focusing (IEF)/SDS–PAGE of cell culture (a) and

hydroponically grown shoot material (b). Sam-

ples before (labeled with Cy3, shown in red) and

after free-flow electrophoresis (FFE)-treatment

(labeled with Cy5, shown in green) were com-

pared. Upper panels: gel pictures of each fluo-

rescence signal. The abundance of spots

encircled in green or black has been decreased

by at least three times as a result of FFE

treatment. Spots encircled in black were picked

for MS/MS analysis. Lower panel: fluorescence

images were electronically overlaid using IMAGE

QUANT TL� software (GE Healthcare, http://

www.gehealthcare.com). Yellow spots represent

proteins of equal abundance before and after

FFE. Spots that are more abundant in the pre-FFE

sample are green, and those more abundant

in the post-FFE sample are red. Arrows indi-

cate proteins unambiguously identified by MS,

the numbers correlate with Tables 1 and 2,

respectively.



new insights into the composition of the mitochondrial

proteome.

In the cell culture differential analysis (Figure 4a, Table 1),

the catalase proteins At1g20620, At1g20630 and At4g35090

were shown to be depleted between 5 and 11 times along

with other peroxisomal proteins, but all three catalases were

reported in Percoll-purified mitochondria (Heazlewood

et al., 2004). Thus, much if not all of the catalase in Percoll-

purified mitochondria is likely to be of peroxisomal origin.

Interestingly, the 3-keto acyl thiolase (At2g33150) in Table 1

had been found in Percoll-purified mitochondria in several

reports (Heazlewood et al., 2004; Kruft et al., 2001), and was

considered a putative dual-targeted protein (Taylor et al.,

2004). But recently a combination of in vitro import assays

and GFP localization studies have provided convincing

evidence for a single location of this protein in peroxisomes

(Carrie et al., 2007), consistent with its greater than 10-times

removal by FFE (Table 1). The triosephosphate isomerase

(At3g55440) in Table 1 was also found in previous studies of

mitochondria, but was convincingly shown to be associated

with the outside of the outer membrane as part of the

association of glycolytic enzymes with mitochondria (Giege

et al., 2003). The greater than 5-times removal of this

cytosolic protein by FFE separation is also supported by

these findings (Table 1). Four other likely plastid contami-

nants that have been previously noted in mitochondrial lists

(Heazlewood et al., 2004) show 7- to 9-times decrease

in abundance (At2g04030, At2g28000, At3g58610 and

At5g13420) in the FFE-purified mitochondria.

In the shoot mitochondrial differential analysis (Figure 4b,

Table 2) a different triosephosphate isomerase was found to

be depleted (At2g21170). This protein had previously been

reported in Percoll-separation mitochondria (Heazlewood

et al., 2004), but is also found extensively in reports on the

plastid proteome (Froehlich et al., 2003; Giacomelli et al.,

2006; Kleffmann et al., 2004; Peltier et al., 2006). Its 9-times

loss by FFE is consistent with the removal of a plastid

contaminant. Three additional likely plastid contaminants

(At2g28000, At5g50920 and At3g48870) that were 4- to

7-times decreased by FFE have also been previously listed

in the mitochondrial proteome (Heazlewood et al., 2004).

The glutamine synthetase GLN2 (At5g35630) has not been

previously claimed by mass spectrometry in mitochondria,

but is claimed to be dual-targeted to mitochondria and

plastids based on GFP localization and antibody studies

(Taira et al., 2004). Table 2 shows this protein was nearly

Table 1 Proteins from cell culture mitochondrial samples are found to vary in average ratio of abundance (ratio >3, P < 0.05) when comparingsamples from before and after free-flow electrophoresis (FFE). Identity of proteins were determined by tandem mass spectrometry (MS), thepredicted molecular weight (MW) and isoelectric point (pI) of the match is shown along with the MOWSE score (P < 0.05 when score >38),number of peptides matched to tandem mass spectra and the percentage coverage of the matched sequence. The putative intracellular locationof these proteins according to a subcellular localization database (SUBA, http://www.suba.bcs.uwa.edu.au) is given to the far right

SpotNo.

AccessionNo.

Averageratio P-value Description MW/pI

MS/MS

Putative cellularlocationScore Peptides

Sequencecoverage

1 At2g04030 6.83 0.037 Heat shock protein 88-1 88663/4.65 640 20 31% Plastid2 At3g60750 5.99 0.015 Transketolase 79968/6.32 452 14 19% Plastid3 At4g24280 6.64 0.020 HSP70-like protein 76508/4.81 127 6 5% Plastid4 At5g49910 7.47 0.015 Heat shock protein 70 76996/4.92 133 5 6% Plastid5 At1g74030 5.98 0.013 Enolase 51474/5.98 193 10 24% Cytosol/nucleus6 At2g28000 7.21 0.018 Chaperonin 60 62072/4.82 836 23 34% Plastid7 At4g35090 10.84 0.010 Catalase 2 56931/7.12 199 10 11% Peroxisome8 At1g20620 5.06 0.016 Catalase 3 56695/7.69 62 4 7% Peroxisome9 At4g20360 7.26 0.029 Elongation factor Tu 51630/6.09 145 6 15% Plastid

10 At3g58610 7.81 0.024 Ketol-acid reductoisomerase 63812/6.79 130 7 8% Plastid11 At4g35090 9.58 0.007 Catalase 2 56931/7.12 87 3 6% Peroxisome12 At1g20630 11.82 0.005 Catalase 1 56762/7.44 164 6 12% Peroxisome13 At2g42790 6.05 0.002 Citrate synthase 56175/7.79 57 4 4% Peroxisome14 At2g33150 11.76 0.005 3-Ketoacyl-CoA thiolase 48579/8.47 131 7 20% Peroxisome15 At5g13420 8.32 0.010 Transaldolase 47699/6.32 105 4 8% Plastid16 At5g16970 5.07 0.036 P1 zeta crystallin 38134/6.10 136 7 16% Cytosol/nucleus17 At3g14420 9.12 0.002 Putative glycolate oxidase 40341/9.58 58 2 4% Peroxisome18 At3g55010 7.81 0.022 Phosphoribosylformylglycinamidine

cyclo-ligase precursor41504/5.14 237 5 13% Plastid

19 At5g19440 5.53 0.046 Putative cinnamyl-alcoholdehydrogenase

35594/7.15 144 8 17% Unclear

20 At3g55440 5.06 0.024 Triosephosphate isomerase 27169/5.17 80 4 19% Cytosol21 At4g05530 8.49 0.003 Putative short-chain

dehydrogenase/reductase26765/8.68 76 4 13% Peroxisome

22 At1g69740 6.78 0.018 Aminolevulinate dehydratase 46690/7.36 118 6 13% Plastid



13-times reduced in abundance by FFE separation, suggest-

ing that most of the protein found in the pre-FFE mitochon-

drial samples is likely to be plastidic in origin and can be

removed along with other plastid contaminants. Addition-

ally, a photosystem II-like chlorophyll a/b binding protein

(At1g29910), which was identified in a proteomic study of

the mitochondrial membrane (Brugiere et al., 2004), is

shown here to be nearly 4-times decreased in abundance

by FFE, and hence is likely to be a plastid contaminant in

mitochondrial samples.

The purity and homogeneity of the cell culture samples

was also assessed by electron microscopy (Figure 5). Pic-

tures A, B and C are taken from material after differential

centrifugation and Percoll density centrifugation, pictures D,

E and F were taken from the same material that has also

been FFE purified. On the pictures, mitochondria can be

recognized as circular structures displaying a network-like

cluster of thick lines inside, which represent the cristae.

Comparing the close-up views of the pre-FFE sample and the

post-FFE sample, a significant loss of particles with a non-

mitochondrial structure is obvious. Therefore, these electron

micrographs clearly demonstrate a gain in the level of purity

of mitochondria prepared from cell culture that is in-line with

the other results presented above.

Integrity of FFE-separated mitochondria

On the electron micrographs in Figure 5, the majority of the

organelles in the FFE-treated sample as well as the control

sample seem to be intact, and no differences between the

two samples in respect to the integrity of the organelles were

detected. This is in contrast to the studies of Zischka et al.

(2003), where a significant higher proportion of the organ-

elles were disrupted in the control sample. However, in that

Table 2 Proteins from shoot mitochondrial samples were found to vary in average ratio of abundance (ratio >3, P < 0.05) when comparingsamples from before and after free-flow electrophoresis (FFE). Identity of proteins were determined by tandem mass spectrometry (MS), thepredicted molecular weight (MW) and isoelectric point (pI) of the match is shown along with the MOWSE score (P < 0.05 when score >38),number of peptide matched to tandem mass spectra and the percentage coverage of the matched sequence. The putative intracellular locationof these proteins according to a subcellular localization database (SUBA, http://www.suba.bcs.uwa.edu.au) is given to the far right

SpotNo. Accession No.

Averageratio P-value Description MW/pI

MS/MS

Putative cellularlocationScore Peptides

Sequencecoverage

1 At5g38410 27.7 0.0002 RuBisCO small subunit 3B 20.3/8.2 113 3 16% PlastidAt1g08480 Succinate dehydrogenase subunit 6 15.8/6.8 73 1 6% Mitochondrion

2 At1g67090 25.7 0.0001 RuBisCO small subunit 1A 20.2/7.8 134 3 20% Plastid3 At3g62030 17.5 0.0002 Peptidylprolyl isomerase ROC4 28.2/8.8 44 2 6% Plastid4 At5g66570 3.5 0.04 Photosystem II oxygen-evolving

complex 33 (OEC33)35.1/5.3 164 5 13% Plastid

5 At2g21170 9.4 0.004 Triose-phosphate isomerase 33.3/8.0 60 1 4% Plastid6 At1g61520 3.9 0.0003 Light-harvesting chlorophyll

a/b binding protein29.2/9.1 55 4 6% Plastid

7 At1g29910 3.8 0.0004 Photosystem II type I chlorophylla/b binding protein, putative

28.2/5.1 75 4 10% Plastid

8 At5g66570 3.1 0.008 Photosystem II oxygen-evolvingcomplex 33 (OEC33)

35.1/5.3 164 5 13% Plastid

9 At2g21330 4 0.04 Fructose bisphosphate aldolase 42.9/6.5 146 6 13% Plastid10 At5g35630 13.1 0.001 Glutamine synthetase 47.4/6.9 38 2 3% Plastid and/or

mitochondrion11 At2g39730 12.9 0.004 RuBisCO activase 52.0/5.9 166 5 12% Plastid12 At2g28000 7.3 0.01 60 kDa Chaperonin alpha subunit 62.0/4.8 518 8 15% Plastid13 At4g24280 10.2 0.004 Heat shock protein 70, putative 76.5/4.8 181 5 5% Plastid14 At3g60750 5.7 0.03 Transketolase-like protein 80.0/6.3 291 13 14% Plastid15 At5g50920 4.3 0.05 ATP-dependent Clp protease

ATP-binding subunit (ClpC1)103.4/6.7 278 10 9% Plastid

At3g48870 ATP-dependent Clp proteaseATP-binding subunit (ClpC2)

105.8/6.3 191 8 7% Plastid

16 At3g60750 4.1 0.008 Transketolase, putative 80.0/6.3 242 7 10% PlastidAt5g08670 ATP synthase beta subunit 59.7/6.6 173 5 8% Mitochondrion

17 AtCg00490 4.4 0.03 Large subunit of riblose-1,5-bisphosphate carboxylase/oxygenase

52.9/6.2 790 23 31% Plastid

18 At3g09260 5.5 0.006 Beta-glucosidase, putative 59.7/6.9 68 2 4% ER19 At3g09260 6.8 0.002 Beta-glucosidase, putative 59.7/6.9 407 17 25% ER20 At5g04590 4.5 0.003 Sulphite reductase 71.9/8.5 84 5 5% Plastid



study, the isolation protocol of the control sample included

only a differential centrifugation step, and did not employ a

Percoll gradient. To further test the integrity of the isolated

mitochondria, individual assessments for the outer and

inner mitochondrial membrane of organelles isolated from

cell culture were performed using an oxygen electrode. In

the presence of cytochrome c and a reductant, the activity of

cytochrome c oxidase before and after the addition of the

detergent Triton X-100 has been used to verify outer mem-

brane integrity (Neuburger et al., 1982). In cell culture sam-

ples the outer membrane integrity of samples ranged from

80–95% pre-FFE to 87–95% post-FFE, whereas from leaf

samples integrity ranged from 94–96% pre-FFE to 95–97%

post-FFE. This demonstrates that FFE treatment did not

damage organelle integrity, and that sometimes it increased

the percentage of mitochondria with intact outer mem-

branes when compared with a sample that has been purified

on a Percoll gradient alone.

Respiratory capacity and inner membrane integrity was

tested by observing oxygen consumption by TCA cycle

substrate, measuring the effect of added ADP on the rate

of oxygen consumption (respiratory control ratio) and

calculating the oxidative phosphorylation efficiency (ADP:O

ratio). Figure 6 shows the respiratory rate of mitochondria

and the effect of ADP additions during succinate- and

malate + glutamate-dependent respiration. Respiratory

rates on a mitochondrial protein basis were always in-

creased following FFE, indicative of removal of damaged

organelles and contaminating proteins (Figure 6). Across all

the samples analysed the respiratory control ratios were 1.3

pre-FFE to 1.4 post-FFE with succinate as a substrate, and 1.9

pre-FFE to 2.0 post-FFE with malate + glutamate as a sub-

strate. The P:O ratios were relatively constant pre-FFE and

post-FFE, at 1.75 for succinate and 2.4 for malate + gluta-

mate, on average. These results show that electron transport

in pre-FFE and post-FFE mitochondrial samples was coupled

to ATP synthesis, a strong indication for the presence of

mitochondria with intact inner membranes.

Organelle yield

The efficiency of a purification step can be expressed as the

ratio of the increase in purity over the loss of sample. A 5- to

6-times reduction in catalase activity has been measured for

(a)

(d)

(b)

(e)

(c)

(f)

Figure 5. Electron micrographs of mitochondrial fractions after differential centrifugation and density gradient centrifugation. Pictures a, b and c were taken from

non-free-flow electrophoresis (non-FFE) treated material. The sample displayed in d, e and f received FFE-treatment. Pictures a and d were taken using a

magnification of ·5000. Pictures b–e (magnification ·15 000) show representative areas of a and c, respectively. M, mitochondrion.



the cell culture mitochondrial extracts, whereas the average

change in abundance of the identified proteins (above the

selected threshold) in the DIGE experiment was a 7-times

reduction. For the shoot mitochondria, an average 9-times

decrease has been calculated based on the average change

of the identified proteins (above the threshold in the DIGE

experiment). In our experience, these values are much

higher than what can be obtained with an additional Percoll

gradient. The loss of sample during the FFE procedure is

approximately 50% (data not shown), which matches the

losses we experienced with Percoll gradients. Based on the

A280 protein quantification shown in Figure 1, the main peak

for the cell culture as well as the shoot material made up

only 60% of the total protein collected from the FFE, with the

residual 40% found in the fractions outside of the main peak.

Limitations of FFE

Although in principle mitochondria preparations for any

purpose can profit from an increased purity, often time

constraints have to be considered in the planning of an

experiment. In ZE-mode, the sample is continuously fed into

the separation chamber. The maximum speed for this is

largely determined by the sample concentration and its

tendency to form aggregates. A slower injection speed will

yield a narrower sample beam in the separation chamber,

which will split into sharper major and minor beams during

the separation process, whereas a faster injection speed will

tend to decrease resolution. FFE can be carried out faster

than a second Percoll gradient for small sample sizes

(1–5 mg of protein), especially if the organelles have to be

washed between the two gradient steps. Large samples

(>10 mg of protein), however, often require more time than a

second Percoll gradient. This is important for samples

requiring fast processing, for example when the effect of

treatment is being assessed, or when long periods of

experimental analysis of the fresh mitochondrial sample are

planned after the isolation procedure.

Experimental procedures

Arabidopsis cell and hydroponic culture

A heterotrophic Arabidopsis cell culture has been maintained underthe same conditions as described in Millar et al. (2001). Conditionsfor the hydroponic culture were adapted from Schlesier et al. (2003).Approximately 50–100 seeds were sterilized and dispensed on astainless steel wire mesh platform (40 mm · 40 mm, elevation20 mm, mesh size 1 mm) layered previously with 1% (w/v) sterilizedagarose in a round plastic vessel (diameter 68 mm, height 76 mm)containing approximately 60 ml of liquid media [¼-strengthMurashige & Skoog medium without vitamins, ¼-strength GamborgB5 vitamins solution, 2 mM 2-(N-morpholine)-ethanesulphonic acid(MES), 1% (w/v) sucrose, pH 5.8). Arabidopsis plants were grownunder long-day conditions with a light intensity of100–125 lmol m)2 sec)1 at 20�C for 21 days. Liquid medium wasreplaced every 7 days.

Mitochondria isolation by differential centrifugation and

Percoll density centrifugation

Isolation of mitochondria from cell culture was carried out accord-ing to the method described by Meyer et al. (2005). Shoot mito-chondria were isolated from 21-day-old Arabidopsis hydroponiccultures by a method developed for pea leaves (Day et al., 1985).Prior to FFE, both enriched mitochondrial fractions were washedthree times with FFE separation medium (10 mM acetic acid, 10 mM

triethanol amine, 1 mM EDTA, 280 mM sucrose) to quantitativelyremove the Percoll.

Free-flow electrophoresis

Free-flow electrophoresis was performed using the BD Free FlowElectrophoresis system (Becton, Dickinson & Company, http://www.bd.com) with a separation chamber height of 0.5 mm. Sepa-ration and counterflow medium (10 mM acetic acid, 10 mM triethanol

(a) (b)

(c) (d)

Figure 6. Assessment of respiratory activity and

integrity of mitochondria before and after free-

flow electrophoresis (FFE) by oxygen electrode

measurements.

(a) Succinate-dependent respiration of cell cul-

ture mitochondria.

(b) Malate + glutamate-dependent respiration of

cell culture mitochondria.

(c) Succinate-dependent respiration of hydro-

ponic shoot mitochondria.

(d) Malate + glutamate-dependent respiration of

hydroponic shoot mitochondria. Rates are given

in nmol O2 min)1 mg)1 protein. Before and after

FFE samples were processed in different oxygen

electrodes runs and overlaid. The arrowheads on

the graph indicate the points at which a certain

substrate or inhibitor was injected to a specified

final concentration: A, ADP (200 lM for cell

culture, 300 lM for shoot material); G, glutamate

(10 mM); K, KCN (1 mM); M, malate (10 mM); S,

succinate (5 mM).



amine, 1 mM EDTA, 280 mM sucrose; media inlets 2–6 and counter-flow inlets 1–3) as well as electrode stabilization medium (100 mM

acetic acid, 100 mM triethanol amine, 10 mM EDTA, 200 mM sucrose;media inlets 1 and 7) were injected into the separation chamber at aspeed of 200 ml h–1. Media for anode and cathode circuits consistedof 100 mM acetic acid, 100 mM triethanol amine and 10 mM EDTA. Avoltage of 600 V was applied, which resulted in a current of�103 mA.Before the FFE run, the sample (�10 lg protein ll–1) was subjected toone stroke in a Potter Elvehjem homogenisator. Sample injectionspeed was 3000–3500 ll h–1 depending on the sample and the levelof contamination. Fractions were collected in 2-ml 96-well plates. Theseparation chamber was cooled to 5ºC, and the sample and 96-wellplates were cooled in an ice bath.

One-dimensional SDS–PAGE and immunoblotting

Precast gels [10–20% (w/v) acrylamide, Tris–HCl, 1 mM were used foranalytical purposes and western blotting. Protein assays (Bradford,1976) were performed for pooled FFE fractions. Proteins were trans-ferred onto nitrocellulose membranes and probed with 1:5000 dilu-tion of the primary antibodies (mtHSP70:PM003 from Dr Tom Elthon,Lincoln, NE, USA; Kat2 (Germain et al., 2001); Rbc, raised againsttobacco Rubisco in rabbits, supplied by Dr Spencer Whitney, Aus-tralian National University). Horseradish-peroxidase-conjugatedsecondary antibodies in a dilution of 1:10 000 were used for thechemiluminescent detection of the immune signal.

DIGE two-dimensional IEF/SDS–PAGE

Samples (50 lg) of pre- and post-FFE, as well as 50 lg of a 1:1 mixtureof both samples, were acetone-precipated, re-solubilized in lysisbuffer [7 M urea, 2 M thiourea, 4% (w/v) CHAPS, 40 mM Tris base,pH 8.5] and individually labeled with 400 lM of weight- and pI-mat-ched fluorescent dyes Cy2, Cy3 and Cy5 (GE Healthcare, http://www.gehealthcare.com), combined and separated on IEF strips(pH 3–10NL, 24 cm; GE Healthcare) according to the manufacturer’sinstructions. After a brief wash in 1x gel buffer, the strips weretransferred on top of a 12% acrylamide glycine gel where they werecovered with 1.2% agarose in 1x gel buffer. Two-dimensional gelswere run at 50 mA per gel for 6 h. Proteins were visualized on aTyphoon� laser scanner (GE Healthcare) and image comparison wasperformed using the DECYDER� software package (version 6.5; GEHealthcare). Three independent experiments were performed andeach of the resulting three gel sets were first analysed using the dif-ferential in-gel analysis (DIA) mode DECYDER prior to a comprehen-sive biological variance analysis (BVA) including all three gel sets. Gelspots were filtered according to their presence (in nine out of nine gelimages), average ratio in abundance (either £ )3 or ‡3 times) andStudent’s t-test value (£0.05). Gel pictures were electronically overlaidusing the IMAGE QUANT TL� software (GE Healthcare).

Electron microscopy

Sample preparation for transmission electron microscopy wasperformed as outlined in Usher et al. (2006).

Oxygen electrode and chlorophyll measurements

A Hansatech Oxygraph oxygen electrode (http://www.hansatech-instruments.com) was used for catalase assays and the determina-tion of inner and outer membrane integrity. All measurements wereperformed in 1 ml of mitochondria reaction buffer [0.3 M sucrose,

5 mM KH2PO4, 10 mM TES (N-tris[Hydroxymethyl]methyl-2-ami-noethanesulfonic acid) 10 mM NaCl, 2 mM MgSO4, 0.1% (w/v) BSA,pH 7.2]. For catalase assays, 4 ll of a 1% (v/v) H2O2 solution wasinjected into the chamber and the rate of O2 evolution was measured.Chlorophyll concentrations of isolated mitochondria were estimatedusing an established method (Arnon, 1949). Absorbance readingswere taken at 645 and 663 nm using an ND-1000 spectrophotometer(NanoDrop Technologies, http://www.nanodrop.com).

Mass spectrometry

Protein samples to be analyzed were cut from the gels and werein-gel digested according to the method described by Taylor et al.(2005). Samples were then dried down in a vacuum centrifuge,resuspended in 5% (v/v) acetonitrile and 0.1% (v/v) formic acid andanalysed on an XCT Ultra IonTrap mass spectrometer according tothe method described by Meyer et al. (2007). Results were queriedagainst an in-house Arabidopsis database comprising ATH1.pep(release 6) from the Arabidopsis Information Resource (TAIR, http://www.arabidopsis.org) and the Arabidopsis mitochondrial andplastid protein sets (database contained a total of 30 700 proteinsequences with 12 656 682 residues) using the MASCOT searchengine version 2.1.04 (Matrix Science, http://www.matrixscience.com) using the tolerances and criteria outlined in Meyer et al. (2007).

Acknowledgements

This work was supported by grants from the Australian ResearchCouncil (ARC) through the Discovery Program and the Centres ofExcellence Program. HE is supported as an ARC Australian Post-doctoral Fellow, AHM as an ARC Australian Professorial Fellow, andCPL is supported by an Australian Postgraduate Award.

Supplementary Material

The following supplementary material is available for this articleonline:Table S1. Details of the identity of proteins found in the 4th peakfound in FFE isolations.This material is available as part of the online article from http://www.blackwell-synergy.com

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Free-flow electrophoresis for purification of plant mitochondria by surface charge

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