- Open Access
Cryo-electron tomography reveals four-membrane architecture of the Plasmodium apicoplast
- Leandro Lemgruber†1,
- Mikhail Kudryashev†2,
- Chaitali Dekiwadia3,
- David T Riglar4, 5,
- Jake Baum4, 5,
- Henning Stahlberg2,
- Stuart A Ralph3Email author and
- Friedrich Frischknecht1Email author
© Lemgruber et al.; licensee BioMed Central Ltd. 2013
Received: 22 November 2012
Accepted: 15 January 2013
Published: 19 January 2013
The apicoplast is a plastid organelle derived from a secondary endosymbiosis, containing biosynthetic pathways essential for the survival of apicomplexan parasites. The Toxoplasma apicoplast clearly possesses four membranes but in related Plasmodium spp. the apicoplast has variably been reported to have either three or four membranes.
Cryo-electron tomography was employed to image merozoites of Plasmodium falciparum and Plasmodium berghei frozen in their near-native state. Three-dimensional reconstructions revealed the number of apicoplast membranes and the association of the apicoplast with other organelles. Routine transmission electron microscopy of parasites preserved by high-pressure freezing followed by freeze substitution techniques was also used to analyse apicoplast morphology.
Cryo-preserved parasites showed clearly four membranes surrounding the apicoplast. A wider gap between the second and third apicoplast membranes was frequently observed. The apicoplast was found in close proximity to the nucleus and to the rhoptries. The apicoplast matrix showed ribosome-sized particles and membranous whorls.
The Plasmodium apicoplast possesses four membranes, as do the apicoplasts of other apicomplexan parasites. This is consistent with a four-membraned secondary endosymbiotic plastid ancestor.
All modern plastids are remnants of a primary endosymbiosis that occurred around one billion years ago, whereby a cyanobacterium was engulfed and retained by a eukaryote. The algae produced by this event possess plastids with two membranes corresponding to the double membranes of the gram-negative cyanobacteria. These algae later diverged into groups known today as Viridiplantae, Glaucophyta and Rhodophyta, and their plastids are known as primary endosymbionts. All existing plastids are either found in these algae, or in later eukaryotes that phagocytosed plastid-bearing organisms and retained their enclosed plastid . Plastids acquired through capture of an organism that contained a primary endosymbiont are referred to as secondary endosymbionts. These organelles would initially have possessed four membranes, deriving from the two primary endosymbiont membranes, plus the plasma membrane of the primary endosymbiont and the phagosomal membrane of the engulfing eukaryote. In some lineages, one of those outer membranes has been secondarily lost to give rise to three-membrane plastids . Algae with secondary endosymbionts can themselves be captured and retained (tertiary endosymbiosis) giving rise to plastids with more than four membranes .
Although all extant plastids are believed to derive from a single primary endosymbiosis event, secondary endosymbiosis has occurred a number of times, giving rise to multiple, unrelated plastid-bearing groups. One such hypothesized group is the chromalveolata, proposed to have arisen from a single secondary endosymbiosis involving a rhodophyte (red alga), that gave rise to extremely diverse organisms such as diatoms, dinoflagellates and apicomplexans .
The plastid of apicomplexan parasites - the apicoplast - has received considerable attention, in part because of the evolutionary implications of the presence of a plastid in this phylum [5, 6], but mainly because apicomplexans are of immense medical and veterinary significance and the apicoplast is the target for important antiparasitic drugs . If the chromalveolate hypothesis holds, the common ancestor of all apicomplexans possessed a four-membrane plastid, although at least one apicomplexan genus, Cryptosporidium, has subsequently lost its plastid. Apicoplasts lack a photosynthetic apparatus, but a recently identified close relative of the Apicomplexa, the alga Chromera, is photosynthetic and possesses a four-membrane plastid that appears to derive from the same red algal origin as the apicoplast .
Descriptions of apicoplast membranes are many and varied. Because the apicoplast identity was established long after apicomplexans had been ultrastructurally investigated there are many descriptions of the apicoplast with differing generic names, eg, spherical body, hohlzylinder and Golgi adjunct. Micrographs since the description of the apicoplast establish plastids with four membranes for many of the apicomplexans, including Toxoplasma gondii, Sarcocystis, Garnia gonadati, and Babesia bovis as do many of the older micrographs for diverse apicomplexans (discussed in ). However, descriptions of the apicoplast of Plasmodium species, the causative agents of malaria, are conflicting. Some preparations appear to support a four-membrane interpretation in sporozoites ; however this was based on a single observation and thus was not further commented upon. In contrast other electron micrographs suggest an apicoplast with only three membranes [16–18]. This has led to an ongoing disagreement in the number of apicoplast membranes of Plasmodium. Loss of one of the four membranes clearly found in other apicomplexans would have considerable molecular consequences for understanding protein trafficking and biogenesis for the Plasmodium apicoplast, so the resolution of this question is desirable.
To investigate this discrepancy, electron microscopy of cryopreserved parasites from Plasmodium falciparum and Plasmodium berghei coupled to tomographic reconstructions was used. Cryo-electron tomography is widely regarded to introduce the fewest artefacts during preparation as the specimen is rapidly frozen (within a few milliseconds) thus preserving molecular details and membrane arrangements [19, 20]. This technique has been used successfully to investigate membranous and cytoskeletal structures in sporozoites of P. berghei[15, 21–23] and Maurer’s clefts of P. falciparum infected red blood cells [24, 25]. Cryo-electron tomography does not include staining with heavy metal salts and thus provides lower contrast, though being sufficient to examine membranes . Reconstructed tomograms of both P. falciparum and P. berghei merozoites clearly show the apicoplast with four delimiting membranes. These membranes often appear paired with a gap between the second and third membranes. In some individual sections through the tomograms, only three membranes are apparent around some apicoplasts, but three-dimensional reconstructions of these resolves the local appositions of two membranes to a total of four delimiting membranes.
All animal experiments were performed according to the FELASA and GV-SOLAS standard guidelines. Animal experiments were approved by the German authorities (Regierungspräsidium Karlsruhe).
Obtaining Plasmodium merozoites
Plasmodium falciparum parasites from strains 3D7 and D10 were maintained using standard procedures. Cultures were grown in human erythrocytes in RPMI 1640 supplemented with L-glutamine, HEPES, hypoxanthine, and gentamycin. Some cultures were supplemented with human heat-inactivated serum and albumax, and others were supplemented with albumax alone. Parasites were incubated at 37°C in a gas mixture of 5% CO2, 1% O2, and 94% N2. For merozoites subject to cryopreservation prior to electron tomography, late schizonts were harvested by magnetic cell sorting and resuspended in medium. Merozoites were mechanically isolated by passage in a needle.
For P. falciparum merozoites to be subject to conventional glutaraldehyde and osmium fixation or for high pressure freezing and freeze substitution (see below), purification was performed as described by Boyle and colleagues .
To obtain P. berghei merozoites, blood of an infected mouse was collected in T-medium (RPMI, 20% FCS, 0.03% gentamicin) plus heparin, centrifuged (110 × g for 8 min) and resuspended in T-medium and placed on a shaker (50 rpm) overnight at 37°C. Mature schizonts were harvested with a Nycodenz gradient (Axis-Shield PoC, Norway), washed and resuspended in RPMI medium prior to freezing.
Tomography was performed essentially as described before [15, 21, 23, 24, 28]. Merozoites in RPMI medium mixed with 10 nm colloidal gold particles were transferred onto glow-discharged holey carbon Quantifoil EM grids. Grids were blotted at 90% humidity using a Vitrobot (FEI). After removing the liquid excess, the material was rapidly plunge frozen into liquid ethane and stored in liquid nitrogen. The grids were observed in a Titan Krios (FEI) operating at 300 kV and equipped with a Gatan post column energy filter and post-GIF CCDs operating at liquid nitrogen temperature. Tilt series were acquired with an increment of 2º covering -60º to +60º, with a cumulative dose under 10,000 electrons/nm2 and a defocus of -3 to -15 μm on a 2k Ultrascan 1000 CCD camera (Gatan).
For this study a total of 12 (for P. berghei) and 24 (for P. falciparum) tomograms were reconstructed by weighted back-projection using the Etomo program in the IMOD software package (Boulder Laboratory for 3D electron microscopy) . Reconstructed tomograms were filtered using non-linear anisotropic diffusion . Visualization, volume rendering and segmentation were performed using the 3dmod program of the IMOD package .
High-pressure freezing and freeze substitution
Purified P. falciparum merozoites from strain D10 were allowed to invade red blood cells, then 10 min post invasion were high-pressure frozen using a LeicaEM high-pressure freezer. The infected erythrocyte samples were freeze substituted with 1% uranyl acetate at -90°C for 24 hrs using a Leica AFS automatic freeze substitution machine. Samples were further freeze substituted with acetone and then infiltrated with increasing concentrations of Lowicryl HM20 resin in acetone. Resin was polymerized using UV light treatment for 48 hrs at -45°C then a further 48 hrs at room temperature. Sections of approximately 90 nanometers were cut at room temperature, stained with uranyl acetate and lead citrate and examined using a Philips CM120 BioTWIN transmission electron microscope at 120 kV (Advanced Microscopy facility, School of Botany, the University of Melbourne, Australia).
Conventional transmission electron microscopy
Purified P. falciparum merozoites (as described above) were fixed in 2.5% glutaraldehyde in pH 7.4 phosphate buffer on ice. Cells were washed in phosphate buffer and fixed in 1% osmium tetroxide (ProSciTech, Australia) in 0.1 M phosphate buffer for one hour on ice. Samples were washed in water, then dehydrated in increasing concentrations of ethanol before embedding in LR Gold resin (ProSciTech, Australia) and polymerized using benzoyl peroxide (SPI-Chem, USA). Sections were cut, stained and inspected as described above.
Results and discussion
The relationship between the apicoplast and the endoplasmic reticulum (ER) has been the subject of some conjecture [16, 31, 34]. In heterokonts the plastid resides within the lumen of the ER , and a similar arrangement is formally possible for the apicoplast (see reviews in [14, 34, 36] for more detailed discussions of such models). Such an arrangement might predict that the outer membrane of the plastid contained ribosomes, necessary for the insertion of apicoplast-targeted proteins. As with previous examinations, such ribosomes on the outer membrane of the plastid, and continuities between the membranes of the plastid and ER were not observed. However, in merozoites, where all organelles are more closely apposed, the apicoplast was observed in close proximity to the nucleus and ER (Figures 3E and 4A), as previously described for Plasmodium, Toxoplasma and Sarcocystis. Although there appeared to be some contact points with the nuclear envelope and with the ER (Figure 3E), no continuity between these membranes is apparent.
Much has been discussed about how proteins are transported across the apicoplast’s surrounding membranes, with various membranous translocon proteins already described [37–41]. In the acquired cryo-tomograms, structures were observed in merozoites’ apicoplast of both P. berghei and P. falciparum, potentially resembling a translocon (Figures 4C and 5C), although this cannot be resolved with the current images. These features were located between the second and the third membranes.
Going through the virtual sections of a reconstructed tomogram, one can observe that depending on the section the apicoplast can show three or four membranes, or even at the same section both arrangements (Figure 5H). This is usually due to the close apposition of the second membrane with the first one, which can explain the discrepancies in the previously published apicoplast structure descriptions [16–18].
The Plasmodium apicoplast was shown to contain four membranes as in other apicomplexan parasites. The apicoplast was found in close proximity to the membranes of the nucleus, inner membrane complex, the mitochondrion and rhoptries. The apicoplast membranes are arranged such that the outer and innermost membranes were closely associated to each other while some space occurred between the two doublet membranes. The presented data reconciles molecular evidence as to how transport into the apicoplast is thought to work in Toxoplasma and Plasmodium with electron microscopy data and shows the capacity of cryo-electron tomography to reveal fine structural details in thin parasites.
We thank Marek Cyrklaff for discussions, Claudia Kuss for P. falciparum culture, Jessica Kehrer and Mirko Singer for help with P. berghei infections and Oliver Grünvogel for helping with the computational modelling.
The work was funded by the Chica and Heinz Schaller Foundation and an ERC starting grant to FF, a postdoctoral fellowship of the University of Heidelberg Cluster of Excellence CellNetworks to LL, the Swiss initiative for Systems Biology (SystemsX.ch, grant CINA) to MK and HS, a Pratt Foundation postgraduate scholarship through the University of Melbourne to DTR. FF is a member of the European Network of Excellence EVIMalaR. DTR received a travel fellowship from the Australian Society for Biochemistry and Molecular Biology and a Harold Mitchell Travel Fellowship, and CD an OzeMalR travel award. SAR is supported by an ARC Future fellowship (FT0990350).
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