Origin, composition, organization and function of theinner membrane complex of Plasmodiumfalciparum gametocytes

Megan K. Dearnley1,2, Jeffrey A. Yeoman3, Eric Hanssen4, Shannon Kenny1,2, Lynne Turnbull5,Cynthia B. Whitchurch5, Leann Tilley1,2 and Matthew W. A. Dixon1,2,*1Department of Biochemistry, Bio21 Institute, 30 Flemmington Road, Melbourne, Vic 3010, Australia2ARC Centre of Excellence for Coherent X-ray Science, The University of Melbourne, Vic 3010, Australia3La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Vic 3086, Australia4Electron Microscopy Unit, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, Parkville, Vic 3010, Australia5The ithree Institute, University of Technology Sydney, NSW 2007, Australia

*Author for correspondence (matthew.dixon@unimelb.edu.au)

Accepted 12 December 2011Journal of Cell Science 125, 2053–2063� 2012. Published by The Company of Biologists Ltddoi: 10.1242/jcs.099002

SummaryThe most virulent of the human malaria parasites, Plasmodium falciparum, undergoes a remarkable morphological transformation as itprepares itself for sexual reproduction and transmission via mosquitoes. Indeed P. falciparum is named for the unique falciform orcrescent shape of the mature sexual stages. Once the metamorphosis is completed, the mature gametocyte releases from sequestration

sites and enters the circulation, thus making it accessible to feeding mosquitoes. Early ultrastructural studies showed that gametocyteelongation is driven by the assembly of a system of flattened cisternal membrane compartments underneath the parasite plasmamembrane and a supporting network of microtubules. Here we describe the molecular composition and origin of the sub-pellicular

membrane complex, and show that it is analogous to the inner membrane complex, an organelle with structural and motor functions thatis well conserved across the apicomplexa. We identify novel crosslinking elements that might help stabilize the inner membranecomplex during gametocyte development. We show that changes in gametocyte morphology are associated with an increase in cellular

deformability and postulate that this enables the gametocytes to circulate in the bloodstream without being detected and removed by themechanical filtering mechanisms in the spleen of the host.

Key words: Gametocyte, Inner membrane complex, Malaria, Plasmodium falciparum

IntroductionMalaria affects more than 240 million people annually and causes

about 781,000 deaths (WHO World Malaria Report 2010; http://

www.who.int/malaria/world_malaria_report_2010/world_malaria_

report_2010.pdf). The apicomplexan parasite Plasmodium

falciparum is responsible for the most deadly form of the disease

in humans. The parasite has a complex lifecycle involving asexual

and sexual reproduction within a human host and an Anopheles

mosquito vector. To prepare itself for life in these different

intracellular niches, the parasite undergoes a remarkable series of

morphological transformations, changing its own shape and

remodelling its host cell environment. The link between asexual

multiplication in the red blood cells (RBCs) of the human host and

sexual reproduction in the midgut of the mosquito is the formation

of a specialized sexual blood stage parasite called the gametocyte.

Late stage P. falciparum gametocytes are characterized by

their unique crescent or falciform shape. Development of these

specialized cells takes at least 7 days and can be divided into five

distinct stages (Baton and Ranford-Cartwright, 2005; Carter

and Miller, 1979; Dixon et al., 2008; Hawking et al., 1971).

Early gametocytes (stage I and IIa) are morphologically

indistinguishable from early asexual parasites. Changes in

cellular architecture are first observed during late stage II to

stage III of development, leading to the characteristic crescent

shape in stage IV, with the gametocyte ends becoming more

rounded in stage V parasites (Dixon et al., 2008; Sinden, 1982;

Sinden et al., 1978).

Like mature asexual stage parasites, stage I–IV gametocytes

sequester away from the peripheral circulation (Day et al., 1998;

Rogers et al., 1996), thereby avoiding passage through and

potential clearance by the spleen. It has been suggested that stage

I–II gametocytes adhere to CD36 before undergoing a switch in

stage III–IV that permits sequestration in the bone marrow,

whereas the mature stage V gametocytes lose the ability to

cytoadhere (Rogers et al., 2000). Indeed, stage V gametocytes

reappear in the peripheral circulation and this is the only stage

that can complete sexual development after ingestion by a

mosquito. Gametocyte maturation thus represents a ‘bottle-neck’

in parasite development; inhibition of this process ablates disease

transmission. Despite the importance of this stage in efforts to

eradicate malaria, relatively little is known of the mechanisms

controlling gametocyte shape, form and function.

Coincident with the adoption of the unique crescent shape of

the late stage gametocyte is the appearance of a tri-laminar

membrane structure known as the sub-pellicular membrane

complex. Early electron microscopy studies on mammalian

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malarial parasites identified this structure as a double-membraned cisternal compartment underlying the parasiteplasma membrane (Sinden and Smalley, 1979). In this work,

we describe some of the molecular components of the sub-pellicular complex and show that the complex is closely relatedto the inner membrane complex (IMC) of the invasive stages of

the parasite.

In the motile (zoite) stages of Plasmodium and otherapicomplexa, the IMC is connected to an actin–myosin motorthat drives cellular invasion through a process called ‘glidingmotility’. The glideosome proteins include myosin-A (MyoA),

myosin-A tail domain interacting protein (MTIP) (Bergman et al.,2003; Herm-Gotz et al., 2002) and the glideosome-associatedproteins 45 and 50 (GAP45 and GAP50) (Gaskins et al., 2004). P.

falciparum GAP50 is an integral membrane protein that anchorspre-complexed GAP45–MTIP–MyoA (Johnson et al., 2007).Host cell invasion is thought to require binding of the actin–

myosin complex to adhesive proteins such as the merozoitethrombospondin-related adhesive protein (mTRAP). mTRAP isembedded in the parasite plasma membrane and binds to

receptors on the host cell, thereby providing the force thatdrives invasion (Baum et al., 2006a). Underlying the IMC is amicrotubule scaffold that forms a major structural element of theinvasive stages of apicomplexan parasites (Kudryashev et al.,

2010; Sinden, 1982).

First, we show that the sub-pellicular membranes ofgametocytes have the same core components as in the invasivestages and that these components are in complex. Second, we

provide evidence that a coordinated recruitment of the IMC to theparasite periphery and assembly of a network of microtubulesdrives elongation of the P. falciparum gametocyte, confirming at

a molecular level suggestions from early electron microscopystudies (Sinden, 1982). This shape change is associated with anincrease in cellular deformability and might explain how latestage gametocytes can survive circulation through the blood

stream of the host.

ResultsIMC proteins are present in complex at the periphery ofmature P. falciparum gametocytes

The late stage P. falciparum gametocyte is characterized by its

unique crescent shape. Coincident with the adoption of this shapeis the appearance of a tri-laminar structure known as thesub-pellicular membrane complex, consisting of a cisternal

compartment underlying the parasite plasma membrane (Sindenand Smalley, 1979). We performed high-resolution serial sectionelectron tomography of this region in a mature stage gametocyte.Fig. 1A shows a virtual section from the tomogram and a

rendered model of the membrane complex. The RBC membrane(red), parasitophorous vacuole membrane (PVM, blue), parasiteplasma membrane (PPM, yellow), the sub-pellicular membranes

(green) and a layer of sub-pellicular microtubules (MT, gray) areevident.

We were struck by the ultrastructural similarities between thissub-pellicular complex (Fig. 1A) and the IMC of merozoites

(Fig. 1B). In the merozoite, the IMC is a cisternal compartmentthat is flattened against the parasite plasma membrane (Fig. 1Bi).It is supported by two or three longitudinally running

microtubules that extend along one side of the merozoite(Fig. 1Bii, arrows), as reported previously (Bannister et al.,2000).

Although previous studies have described the sub-pellicular

membrane complex of the P. falciparum gametocyte at the

ultrastructural level, there are few reports on the composition or

origin of this organelle. In an effort to determine whether the

gametocyte membrane structure is indeed related to the IMC, we

performed fixed cell immunofluorescence microscopy using

antibodies recognizing known components of the merozoite

IMC. Fig. 2A–D shows the presence of GAP50, GAP45, MTIP

and MyoA at the periphery of stage IV gametocytes, consistent

with a location in the sub-pellicular membrane complex. At this

level of resolution, the profile was similar to that of the P.

falciparum PVM protein, Pfs16, which has been shown to be

gametocyte-specific (Baker et al., 1995). Protein solubility and

expression profiling by western blotting of gametocytes and

schizont stage parasites confirmed the presence of the IMC

components (supplementary material Fig. S1, and below).

We have previously generated transfectants expressing a GFP

chimera of P. falciparum GAP50 as a tool to assess IMC genesis

and reorganization during the development of merozoites

(Yeoman et al., 2011). We transfected the P. falciparum

GAP50–GFP gene construct into a high gametocyte producing

clone of 3D7 and found that the GAP50–GFP chimera is

correctly expressed and present at the periphery of mature stage

gametocytes, co-locating with endogenous GAP45 (Fig. 3A). To

determine whether components of the IMC are in complex,

we performed immunoprecipitation experiments. Stage III–IV

gametocytes expressing the GAP50–GFP chimera were

magnet-enriched and then solubilized using RIPA detergent.

The cleared lysate was incubated with rabbit antibodies against

P. falciparum GAP45, and immunoprecipitated using Protein A/

G Sepharose. Precipitated samples were separated by SDS-PAGE

Fig. 1. Trilaminar complex of the gametocyte is closely related to the

inner membrane complex of invasive stages of P. falciparum. (A) Electron

tomogram showing the membrane and microtubule organization in a stage IV

gametocyte. (Ai) Selected virtual tomogram section (22 nm). (Aii) Rendered

model generated from four serial sections through the region shown in Ai.

The rendered model shows the RBC membrane (red), PVM (blue), PPM

(yellow), IMC (green) and microtubules (gray). Osmophillic bodies are

represented in gold. (B) Stained thin section through a dividing schizont

showing a nascent merozoite (Bi) and a selected virtual section (7 nm) from a

tomogram of an individual merozoite (Bii). Rhoptry (R), micronemes (m),

apicoplast (a), nucleus (N), microtubule (MT). Scale bars: 100 nm.

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and transferred for western blotting. Blots were probed with anti-

mouse GFP, GAP50 or MTIP. Full-length GAP50–GFP (64 kDa)

was detected in the precipitated pellet of the transgenic sample,

by both the anti-GFP and anti-GAP50 reagents, but not in the

3D7 parent line (Fig. 3B). MTIP (28 kDa) and endogenous

GAP50 (42 kDa) were detected in the precipitated pellets of

both the transfectant and wild-type samples (Fig. 3B). This

demonstrates that both GAP50–GFP and endogenous GAP50 are

in complex with endogenous GAP45 and MTIP. These studies

validate the use of P. falciparum GAP50–GFP as a marker of the

gametocyte IMC.

Western blotting analyses revealed the presence of GAP45,

GAP50 and MTIP from stage II to stage V gametocytes

(supplementary material Fig. S1B) and immunofluorescence

microscopy confirmed that these proteins were present at the

gametocyte periphery (supplementary material Fig. S2A–D).

Moreover, the solubility profiles for these proteins were similar

in gametocytes and merozoites (supplementary material Fig. S1).

These data, in conjunction with the immunoprecipitation data,

show that the gametocyte sub-pellicular membrane complex is

very closely related to the merozoite IMC and we suggest that it

be renamed the gametocyte IMC.

GAP50–GFP is present in the endoplasmic reticulum ofearly stage gametocytes prior to recruitment to theparasite periphery

P. falciparum GAP50 is recruited from the endoplasmic

reticulum (ER) to the nascent IMC during asexual schizogony

(Yeoman et al., 2011). To determine the origin and

reorganization of proteins during the formation of the

gametocyte IMC we examined the location of GAP50–GFP inlive gametocytes that were co-labelled with the membrane probe,

BODIPY-TR-ceramide. During the initial stages of elongation(stage III), when the gametocyte first becomes morphologicallydistinguishable, GAP50 was associated with a reticular structurein the parasite as well as along one side of the elongating

parasite (Fig. 4Ai). Fixed cell immunofluorescence microscopyconfirmed that GAP50–GFP labels internal structures that werealso recognized by an antibody against the P. falciparum ER

resident protein, ERC (ER calcium binding protein) (Fig. 3C). Bycontrast, ERC was absent from the peripheral location to whichGAP50–GFP is recruited as the gametocyte elongates. 3D-

structured illumination microscopy (3D-SIM) can provideenhanced resolution of cellular structures (Schermelleh et al.,2008). Using 3D-SIM we found that the GAP50–GFP-labelledIMC remained closely associated with the intracellular reticular

structure (Fig. 3D,E, arrows). A rendered model of the GAP50–GFP fluorescence illustrates the points of associations (Fig. 3F,arrows), as do translational and rotation views of the fluorescence

micrographs (supplementary material Movies 1, 2). This isconsistent with an early ultrastructural study (Sinden et al.,1978) that reported connectivity between the ER and the sub-

pellicular membranes.

Stage III gametocytes undergo a dramatic cellularrearrangement, adopting a ‘hat-like’ appearance. During this

stage, the GAP50–GFP is concentrated in the flattened rim regionof the ‘hat’, with additional looping structures around the top ofthe cell (Fig. 4Ai,ii; arrows; see supplementary materialMovie 2). As the IMC develops further it adopts a cupped

shape along the foot, in addition to the looping structures.Electron microscopy confirmed the presence of the IMC alongthe flattened surface and around the ‘pinching’ ends of the

gametocyte (Fig. 4Bi,ii). Fig. 4Bi depicts a longitudinal slicethrough a region of the sub-pellicular microtubule layer, with theIMC visible at the periphery. Some of the microtubules that pass

through the plane of the section are indicated with arrows.Fig. 4Bii shows a cross-section in which the microtubules areviewed end-on and the IMC is evident as a layer outside themicrotubule ‘basket’.

In stage IV of parasite development, the characteristic pointedends of the P. falciparum gametocyte become apparent. By thisstage, the GAP50–GFP extends further around the parasite, the

surface of which was delineated by BODIPY-TR-ceramidelabelling (Fig. 4Aiii; see supplementary material Movie 2 for a3D rotation). 3D-SIM imaging of GAP50–GFP revealed a

periodic banding pattern running perpendicular to the directionof gametocyte extension (Fig. 5A, arrows). The average width ofthe GAP50–GFP fluorescence bands was 460640 nm, with the

interleaved regions of low fluorescence being 120640 nm inwidth (Fig. 5A; supplementary material Movie 3).

Cryo-electron tomography allows 3D visualization ofspecimens after plunge-freezing in cryogenic liquids to

preserve membrane structures (Cyrklaff et al., 2007;Kudryashev et al., 2010). Tomographic analysis of a stage IVgametocyte permits visualization of virtual sections near the

surface of the cell without the need for physical sectioning.Fig. 5Bi highlights the microtubule network running underneaththe cell surface and the locations of the IMC and parasite

membranes within the cell. The section in Fig. 5Bii wasphysically located 100 nm above the section in Fig. 5Bi andshows regions of membrane (approximately 400 nm in width)

Fig. 2. Components of the glideosome and IMC are present at the

gametocyte periphery. (A–D) Immunofluorescence microscopy of 3D7

parasites labelled with antibodies against Plasmodium falciparum GAP50,

Plasmodium falciparum GAP45, MTIP or MyoA (green) and the gametocyte

marker, Pfs16 (red). Bright field (BF) images are shown on the right. Scale

bars: 5 mm.

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interleaved with connecting bands (about 100 nm in width),

layered across the underlying microtubules. A higher

magnification view of the microtubules (Fig. 5C) highlights the

spacing and arrangement of the microtubule network within the

mature stage parasite; the microtubules (25 nm diameter) are

spaced at intervals of approximately 10 nm. The spacing, shape

and orientation of the flattened membrane structures are

reminiscent of the GAP50–GFP-labelled features (Fig. 5A;

supplementary material Fig. S3; Movie 5a,b). The intervening

bands might represent areas of deposition of proteins that

stabilize the IMC and the underlying microtubule network.

In stage V, the ends of the gametocytes become rounded

(Fig. 4Aiv, Fig. 5Aiii). This is probably due to dismantling of the

supporting microtubule network as described below and in a

previous study (Sinden et al., 1978). GAP50–GFP persists at the

periphery of the parasite still co-locating with BODIPY-TR-

ceramide (Fig. 4Aiv). 3D-SIM revealed the maintenance of the

IMC around the cell periphery, and the persistence of the stripes

(Fig. 5Aiii, arrows). The gametocyte presented in supplementary

material Movie 3 shows the transition from stage IV to stage V

with one rounded and one pointed end. In this rotation, the stripes

are clearly observed and there is evidence for some remnant

GAP50–GFP fluorescence around the parasite nucleus.

Temporal and spatial map of gametocyte IMC and

microtubules at different stages of gametocyte

development

We investigated the assembly of microtubules during gametocyte

development. The microtubule-labelling reagent, Tubulin

Tracker (Invitrogen) started to accumulate in a peripheral

region of a stage II gametocyte (Fig. 6Ai; supplementary

material Movie 4) before becoming concentrated along one

edge of the stage III gametocyte (Fig. 6Aii; supplementary

material Movie 4). Extension and elaboration of the microtubule

Fig. 3. Plasmodium falciparum GAP50–GFP is trafficked via the ER to the gametocyte IMC and is in complex with known IMC components.

(A) Immunofluorescence microscopy of GAP50–GFP-expressing gametocytes labelled with antibodies against GFP (green) and Plasmodium falciparum GAP45

(red). Bright field (BF) images are on the right. (B) Proteins were detergent-solubilized from enriched, mature stage GAP50–GFP-transfected or parent 3D7

gametocytes. The extract was immunoprecipitated (IP) using rabbit anti-GAP45. Western blots (WB) of the pellet fractions were probed with mouse anti-GFP, anti-

GAP50 and anti-MTIP. WT, wild-type. Molecular mass is shown in kDa. (C–E) Immunofluorescence (C) and 3D-SIM (D,E) of GAP50–GFP transfectants labelled

with anti-GFP (green) and anti-ERC (red). (D) Projection of sections 10–15 (from a 20-section series) and (E) individual sections. DAPI fluorescence is shown on the

right. In C, an accumulation of GAP50–GFP fluorescence can be seen along one side of the stage III parasite. (F) Rendered model of the 3D reconstruction. D, E and F

illustrate the association of the GAP50–GFP-labelled reticular and peripherally located compartments. The single arrows indicate a region of close association

between the ER and the IMC; double arrows show a region of IMC looping that appears to be continuous with the ER compartment. Scale bars: 3 mm.

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network was observed in stage IV gametocytes (Fig. 6Aiii;

supplementary material Movie 4), adopting a pattern analogous

to that of the peripheral population of GAP50–GFP (Fig. 4Aii).

No microtubule filament bundles were observed in the stage V

gametocyte, consistent with the disassembly of the network in

mature gametocytes.

We used the enhanced resolution of 3D-SIM to investigate the

locations of the microtubules and IMC during gametocyte

elongation. Antibody against b-tubulin labelled microtubules

within the ‘foot’ of the developing gametocyte (Fig. 6B, white

arrows) in addition to a diffuse cytoplasmic staining. b-tubulin in

the foot region was co-located with GAP50–GFP (Fig. 6B). The

distribution of the microtubules and the IMC can best be

appreciated in supplementary material Movie 5a,b, which shows

a translational through and a rotation of the same parasite.

GAP50–GFP and b-tubulin were also co-located in the loop

around the top of the gametocyte (Fig. 6Bii; supplementarymaterial Movie 5a,b). Some GAP50–GFP-labelled regionsaround the edge of the foot did not have associated

microtubules (supplementary material Fig. S3; Movie 5a,b).Electron micrographs of equivalent stages show areas wherethe IMC had started to expand but without any underlying

microtubules (Fig. 4Bii, red arrows). By stage IV there was acomplete network of microtubules underlying the peripherallylocated IMC and forming a similar pattern to that for GAP50.

This characteristic shape and the distribution of the microtubulescan be best appreciated in the 3D-SIM reconstruction shown insupplementary material Movie 6.

During stage III we saw a marked difference in the locations ofGAP45 compared with GAP50–GFP and b-tubulin. This protein

was restricted to the foot and was not co-located with the ER-associated population of GAP50–GFP (Fig. 6B,C; supplementarymaterial Fig. S2B).

Gametocyte elongation is not driven by an actin–myosinmotor

Gametocytes do not display gliding motility and it is unlikely thatthe IMC and associated MyoA participate in a motor complex

analogous to that operating in the zoite stages of P. falciparum.To test for a potential role of actin, we examined the effect ofcytochalasin D on gametocyte morphology. Previous work hasshown that merozoites treated with cytochalasin D (0.1–2 mM)

are rendered immobile and unable to invade erythrocytes (Milleret al., 1979; Srinivasan et al., 2011). By contrast, we found thatmaintenance of gametocytes over a period of 6 days in the

presence of cytochalasin D (at concentrations up to 10 mM)did not prevent the morphological changes that accompanydevelopment to stage IV (data not shown). This indicates that

although MyoA interactions might play an important structuralrole, morphological changes are not driven by an actin–myosinmotor.

Gametocyte shape change is associated with alteredcellular deformability

In an effort to determine a functional role for the remarkable shapechanges of gametocytes, we used ektocytometry to measurecellular deformability at different stages of development (Fig. 7).

The elongation index (EI) of synchronous enriched gametocyteswas measured at sheer stresses of 0–20 Pa. Asexual trophozoitestage parasites showed a significant decrease in ability to elongate

under flow (EI50.13) compared to uninfected RBCs (EI50.34) atthe physiologically relevant sheer stress of 3 Pa (Fig. 7A). StageIII gametocytes showed a similarly low deformability (EI50.12)(Fig. 7A). By contrast, there was an increase in the EI upon

transition to stage IV of gametocyte development (EI50.16); thisvalue was significantly higher than that for stage III (P50.0001).A further increase in the EI was observed upon transition to stage

V (EI50.18) (Fig. 7A).

We made a rough quantification of the change in the shape atdifferent stages of development by monitoring length and widthin DIC images of live gametocytes. Mature trophozoite stage

parasites displayed a roughly circular profile with a mean lengthof 4.3160.05 mm and mean width of 4.060.2 mm (Fig. 7B,C).The mean length of the gametocytes increased significantly from

stage III (6.660.4 mm) to stage IV (10.960.4 mm) (P50.0001),with a slight relaxation upon transition to stage V (9.160.4 mm)(P50.005) (Fig. 7B,C). The width of the stage III gametocyte

Fig. 4. IMC is assembled under the gametocyte plasma membrane.

(A) Live cell confocal imaging of stage III (Ai,Aii), stage IV (Aiii) and stage

V (Aiv) gametocytes expressing Plasmodium falciparum GAP50–GFP

(green) co-labelled with BODIPY-TR-ceramide (red). White arrows indicate

an accumulation of GAP50–GFP along the rim of the stage III parasite (Ai)

and a loop around the top of the gametocyte (Aii). Bright field (BF) images

are on the right. 3D rotations of these cells are shown in supplementary

material Movie 2. Scale bars: 3 mm. (B) Electron micrographs of stage III

gametocytes. (Bi) Longitudinal and (Bii) cross-sections showing the RBC

membrane, PVM, PPM and the double membrane of the IMC, above a row of

microtubules (MT). The electron-lucent area probably represents a vacuole

within the gametocyte cytoplasm. Red arrows mark an area where the IMC

has formed but no microtubules are present. Scale bars: 200 nm. See also

supplementary material Fig. S2.

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(3.060.7 mm) was significantly less (P50.0001) than that of the

asexual stage parasite, with a further significant decrease in thestage IV gametocyte (2.660.1 mm, P50.006), before remainingstable for the remainder of its development (stage V;

2.660.2 mm) (Fig. 7B,C). The length-to-width ratio increasedfrom 1.1 (trophozoite) to 2.2 (stage III) to 4.3 (stage IV), and thendecreased to 3.5 (stage V).

DiscussionIn 1880, Alphonse Laveran observed gametocytes in the bloodsmear of an Algerian malaria patient and remarked upon the

crescent shape of the parasite (Laveran, 1880). Over a centurylater we are still battling to understand the molecular basis forthis unusual shape and its role in P. falciparum transmission and

virulence. Early ultrastructural analyses revealed that the grossmorphological restructuring of the gametocyte is coincident withthe appearance of a tri-laminar membrane structure subtended bya layer of structural microtubules (Aikawa and Beaudoin, 1969;

Sinden, 1982; Sinden et al., 1978). Comprising a double-membrane-bound compartment underneath the parasite plasmamembrane, this structure was named the sub-pellicular membrane

complex (Sinden and Smalley, 1979). Indeed the presence ofalveolar sacks subtending the plasma membrane is a unifyingmorphological feature of apicomplexa, dinoflagellates and

ciliates. Different organism groups have adapted this membranestructure for different cellular functions, with roles ranging fromshape maintenance (free-living protozoa), to protective armor

(dinoflagellates) to calcium stores (ciliates). In the motile stagesof apicomplexan parasites, the sub-pellicular membrane is termedthe IMC and plays a vital role as the scaffold for the motor

complex that drives parasite motility (Beck et al., 2010; Frenal

et al., 2010; Gaskins et al., 2004).

Our work confirms previous studies showing that thegametocyte sub-pellicular membrane complex has ultrastructural

similarity to the IMC of the motile stages of P. falciparum

(Bannister et al., 2000; Sinden and Smalley, 1979); however, themolecular similarity between these structures has not previouslybeen investigated. In this work, we show that key proteins of the

merozoite glideosome complex (GAP50, GAP45, MyoA andMTIP) are present in the gametocyte sub-pellicular membranecomplex. We also show that exogenous GAP50–GFP forms a

complex with endogenous GAP45, GAP50 and MTIP. These dataindicate that the gametocyte membrane complex is closely relatedto the IMC of the motile stages of P. falciparum.

An interesting issue is the roles that the IMC and the associatedglideosome proteins play in gametocyte assembly. Gametocytesdo not display gliding motility, suggesting that the MyoA doesnot actively participate in an actin–myosin motor complex.

Furthermore, we showed that treatment with cytochalasin D didnot prevent gametocyte shape change. This indicates that theMyoA interaction with glideosome proteins probably plays a

structural role rather than driving elongation per se. It seemslikely that the microtubular cytoskeleton is attached to (andorganized by) the IMC, which in turn is linked to the PPM via

glideosome proteins. Thus, our data confirm and support theoriginal suggestion (Sinden, 1982) that gametocyte elongation isdriven by the assembly of microtubules underneath the IMC.

The shape adopted by P. falciparum gametocytes isreminiscent of the elongated sporozoite and ookinete. Indeed,the morphological changes that accompany maturation from

Fig. 5. Gametocyte IMC comprises

rectangular membrane plates. (A) 3D-SIM

of stage IV (Ai,ii) and stage V (Aiii)

gamocytes transfected with Plasmodium

falciparum GAP50–GFP showing vertical

striations (white arrows). Rotations of the 3D-

SIM data are presented in supplementary

material Movie 3. The cells are co-stained

with DAPI (blue). Aii is a zoomed view of the

cell in Ai, highlighting the periodic striations.

Scale bars: 3 mm. (B) Two 30 nm virtual

sections from a cryo-electron tomogram of a

region near the surface of a gametocyte. The

section shown in Bii is 100 nm above the

section in Bi. The IMC, parasite membrane

(PM) and microtubules (MTs) are indicated.

The IMC is segmented by vertical striations

of amorphous material (Bii), marked as

‘sutures’. (C) Lying directly underneath, and

running perpendicular to the rectangular

sections of IMC, is a network of

microtubules. Shown is a virtual section of a

cryo-electron tomogram of a longitudinal

section of microtubules. The same section is

shown below rotated 90˚ to obtain a cross-

sectional view. Microtubule singlets (s) of

25 nm in diameter and doublets (d) of 42 nm

are indicated. The microtubules are separated

by a gap (g) of about 10 nm. Scale bars:

100 nm.

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stage II to stage IV of gametocytogenesis resemble (in reverse)

the shape changes that accompany the transformation of

the elongated sporozoites into liver stage trophozoites

(Jayabalasingham et al., 2010) or ookinetes into oocysts (Carter

et al., 2007). Taken together these data suggest that the IMC

plays an important structural role in each of the cellular

remodelling events that drive parasite elongation. This is

consistent with recent work showing that specific IMC proteins

can play roles in both motility and cell morphology (Tremp and

Dessens, 2011).

Recent work from our laboratory indicates that the merozoite

IMC is formed by the redistribution of a sub-compartment of the

ER to nascent apical caps (Yeoman et al., 2011). Similarly, we

show here that P. falciparum GAP50 is located in the ER in early

stage gametocytes and that part of this population is recruited to

the periphery to form the IMC. This is in agreement with early

ultrastructural study of P. falciparum (Sinden et al., 1978) and

with studies from other alveolates (Gould et al., 2008).

Recruitment of GAP50 to the gametocyte periphery appears to

be coordinated with the laying down of microtubules. Because

the microtubule-supported IMC initially develops along one side

of the gametocyte, stage III gametocytes have a ‘hat-like’

appearance. The IMC and the microtubule network then expand

around the periphery, resulting in a cupped leaf shape. Our

electron and fluorescence microscopy data suggest that the IMC

extends first, followed by microtubule deposition, such that the

leading edges of the nascent IMC are outside the supporting layer

of microtubules. Other IMC proteins, GAP45 and MTIP appear

to be recruited onto the nascent IMC. This process might be

similar to the recruitment of pre-complexed GAP45–MTIP–

MyoA onto membrane-embedded GAP50 during the formation

of the Toxoplasma IMC (Gaskins et al., 2004).

3D-SIM imaging of the GAP50–GFP-labelled IMC revealed a

remarkable architectural feature – bands of fluorescence of

,400 nm interrupted by ,100 nm regions of low fluorescence.

Cryo-electron tomography revealed the ultrastructural analogue

of these bands as flattened regions of membrane with intervening

amorphous material. These structures are probably equivalent to

the transverse ‘sutures’ encircling the gametocyte that were

identified in an early freeze-fracture study (Meszoely et al.,

1987). Similar patchworks of sub-pellicular membrane have been

described in the IMC of the invasive stages of P. falciparum, as

well as in Toxoplasma and other apicomplexa (Bannister et al.,

2000; Raibaud et al., 2001). Another early transmission electron

microscopy study identified protein ‘crosslinks’ running

perpendicular to the microtubule network (Kaidoh et al., 1993).

We suggest that the GAP50–GFP-labelled IMC ‘stripes’

represent IMC cisternae laid down as rectangular plates,

whereas the gaps represent proteinaceous material that holds

the cisternae and the microtubules in place.

A concurrent large-scale analysis of IMC components in P.

falciparum revealed a novel IMC protein that appears to be

restricted to the genus Plasmodium (Mal13P1.228). In the early

gametocyte, this protein is located in thin stripes along the ‘foot’,

which expand and eventually encircle the mature stage gametocyte

(Kono et al., 2012). These structures might represent the regions of

low fluorescence and proteinaceous crosslinks identified in our

3D-SIM and cryo-electron microscopy studies.

Consistent with previous ultrastructural studies of gametocytes

(Sinden, 1982; Sinden et al., 1978), there is no morphologically

identifiable apical polar ring in gametocytes to organize the

peripheral microtubules [although a microtubule organizing

center is formed very quickly after activation (Aikawa, 1977)].

This begs the question: how are the microtubules organized

onto the well-ordered cytoskeletal basket? We suggest that

the proteinaceous ‘sutures’ between the IMC plates provide a

means of stabilizing the microtubule network. Proteins resident in

these sutures might function in the targeting of the nascent IMC

Fig. 6. Coordinated development of the IMC and the microtubule

network drives shape change. (A) Live cell confocal imaging of stage II–V

gametocytes (Ai–Aiv) labelled with Tubulin Tracker (TT, green) and

BODIPY-TR-ceramide (red). DIC images are shown on the right. (B) Single

sections from a 3D-SIM reconstruction of stage III Plasmodium falciparum

GAP50–GFP gametocytes prepared for immunofluorescence microscopy

(anti-GFP, green; anti-b-tubulin, red). The white arrows mark areas of

peripheral GAP50–GFP staining with associated b-tubulin. The red arrow in

Bii highlights the looping extensions of the IMC with concentrated b-tubulin

staining. A montage of projections of Bi is shown in supplementary material

Fig. S3. Projections and 3D rotations of Bi and Bii are presented in

supplementary material Movie 5a,b. (C) Stage III gametocyte labelled with

anti-GAP45 (green) and anti-b-tubulin (red). This image highlights the

restricted location of GAP45 within the foot of the gametocyte. Scale bars:

3 mm. See also supplementary material Movies 6,7.

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to the periphery of the cell and in the stabilization of the IMC and

the sub-IMC microtubule network.

Analysis of DIC images revealed a significant increase in the

length and a reduction in the width of the gametocyte during

maturation as compared with asexual stage parasites. A more

detailed analysis of gametocyte shape by cryo-X-ray tomography

gives similar results (Hanssen et al., 2011). The elongated form

of P. falciparum gametocytes is unusual. Gametocytes from other

human malaria species and gametocytes from most other species

of Plasmodium have a more rounded morphology, although

gametocytes of avian plasmodia are also elongate (Sinden et al.,

1978). P. falciparum is also unusual among human malaria

parasites in that late stage asexual parasites drastically remodel

the RBC through the export and insertion of parasite-derived

proteins into and under the host RBC membrane (Tilley et al.,

2011). These modifications lead to a significant increase in

Fig. 7. Gametocyte shape changes are

accompanied by changes in

deformability. (A) Elongation index (EI)

values for uninfected RBCs, asexual

trophozoites (30–36 hours after invasion)

and stage III–V gametocytes were

measured using a RheoScan slit

ektocytometer at shear stresses of 0–20 Pa.

The data represent the mean6s.e.m. of

triplicate measurements and are typical of

experiments performed on two separate

occasions. The red box highlights the

measurements at 3 Pa. (B) Illustrations of

stage III–V gametocytes (female). The

dimensions were measured from DIC

images (mean6s.e.m.). (C) Length-to-

width ratio6s.e.m. for different

developmental stages. a, apicoplast; Ac,

acidocalcisomes; C, cytostome; DV,

digestive vacuole; g, Golgi complex; IMC,

inner membrane complex; k, knob; Lb,

Laveran’s bib; m, mitochondrion; MC,

Maurer’s cleft; Mt, microtubules; N,

nucleus; Ob, osmophillic bodies; PV,

parasitophorous vacuole; RBC, red blood

cell; W, whorl.

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cellular rigidity, which would make mature parasite-infected

RBCs vulnerable to recognition and clearance within the spleen

(Glenister et al., 2002; Safeukui et al., 2008). The parasite

avoids these clearance mechanisms by cytoadhereing within

the microvasculature of the host. Early stage P. falciparum

gametocytes also sequester; however, upon reaching sexual

maturity they release from their sites of adhesion and re-enter the

circulation to enable transmission to mosquitoes (Day et al.,

1998; Hayward et al., 1999).

In an effort to determine the functional consequences of the

gametocyte shape changes we investigated the ability of asexual

trophozoites and stage III–V gametocytes to elongate in flow, a

surrogate measure of cellular deformability (Groner et al., 1980;

Hardeman et al., 1987). In agreement with previous reports, we

found that asexual trophozoite-infected RBCs are highly

undeformable (Nash et al., 1989). This loss of deformability is

probably due to a combination of membrane rigidification and

the presence of the large rigid intracellular parasite (Glenister

et al., 2002; Nash et al., 1989). We found that stage III

gametocytes have similar deformability properties to trophozoite-

infected RBCs; however, upon transition to stage IV the

gametocytes show a significant increase in the elongation index

measured under flow conditions. This is probably due in part to

the elongated shape. A further increase in the elongation index in

stage V gametocytes might be due to the fact that the microtubule

network is disassembled at this point, leaving an already

elongated cell that probably extends further under flow pressure.

These changes in rheological properties might help stage V

gametocytes survive in the circulation of the host. Indeed, stage

V gametocytes might have rheological properties more akin to P.

vivax-infected RBCs (Handayani et al., 2009; Suwanarusk et al.,

2004). As its name suggests (vivax means lively), P. vivax is

more amoeboid than P. falciparum and appears to avoid splenic

clearance in spite of an inability to cytoadhere. Stage III–V

gametocytes lack knobs and are thought to adhere with lower

affinity than asexual stages and to prefer sites of reduced vascular

flow (Rogers et al., 1996; Smalley et al., 1981). It is possible that

another consequence of the elongated shape of P. falciparum

gametocytes is enhanced adhesion to lower affinity receptors by

permitting flattening of the cell, which might increase the area in

contact with the capillary walls.

In conclusion, our data show that the falciform shape of P.

falciparum gametocytes is supported by an IMC with very similar

composition to the merozoite IMC, linked to underlying

microtubules. A previously unrecognized correlate of gametocyte

elongation is an enhanced cellular deformability. We anticipate that

disruption of the IMC and its associated microtubules compromises

the ability of P. falciparum to adopt the falciform shape; this might in

turn affect its ability to survive in the circulation. This could provide

a new avenue for interfering with this important human pathogen.

Materials and MethodsParasite culture and gametocyte enrichment

Parasite-infected RBCs were cultured in RPMI-HEPES supplemented with 5%human serum and 0.25% AlbuMAX II as previously described (Foley et al., 1994).Laboratory strains of 3D7 cultured for long periods form gametocytes with lowefficiency, therefore we used a 3D7 isolate recovered from a patient (Lawrenceet al., 2000) and cultured for only a limited time. A GAP50–GFP transgenicparasite was created by transfecting a high gametocyte producing clone of 3D7with the pGLUX–GAP50–GFP construct as previously described (Yeoman et al.,2011).

Gametocytes were prepared using a modification of a published protocol(Fivelman et al., 2007). A culture of mainly ring stage parasites (6–8%

parasitemia) was treated with 5% sorbitol (Lambros and Vanderberg, 1979),then separated using a Percoll density gradient (Knight and Sinden, 1982) to enrichthe ring stage and remove any gametocytes. The parasites were cultured until theyreached 8–10% trophozoites then sub-divided into 2% trophozoites (5%hematocrit). The culture was maintained for 10 days in the presence of62.5 mM N-acetyl glucosamine to inhibit merozoite invasion and thus asexualreplication (Hadley et al., 1986). Giemsa-stained slides were used to monitor stageprogression. The medium was changed daily but no fresh erythrocytes were added.At the desired stages of development, gametocytes were enriched by magneticseparation (Fivelman et al., 2007). For analysis of the effect of cytochalasin D(Sigma-Aldrich), the drug was added at concentrations of 0–10 mM at day one ofgametocyte generation and supplemented daily into the culture media. Theparasites were assessed by microscopy at different stages of development.

Microscopy and image analysis

Live infected RBCs were prepared for fluorescence microscopy as previouslydescribed (Dixon et al., 2011). Immunofluorescence was performed on acetone-fixed blood films (Spielmann et al., 2006) using the following primary antibodiesin PBS containing 3% BSA: mouse anti-GFP (1:500, Roche), rabbit anti-GFP(1:500), rabbit anti-GAP45 [1:500 (Baum et al., 2006b)], rabbit anti-GAP50 [1:500(Baum et al., 2006b)], rabbit anti-Pfs16 [1:1000 (Baker et al., 1995)], mouse anti-Pfs16 [1:1000 (Dixon et al., 2009)], mouse anti-MTIP [1:500 (Green et al., 2006)],mouse anti-MyoA (Baum et al., 2006b) followed by anti-mouse or anti-rabbit IgGconjugated to Alexa Fluor 568, Alexa Fluor 647 or FITC. Slides were labelled with1 mg/ml of DAPI prior to mounting.

BODIPY labelling of parasite-infected cells was performed by adding BODIPY-TR-ceramide at a final concentration of 0.7 mM to the parasite culture. Cells wereincubated overnight under standard culture conditions. Labelling with TubulinTracker Green was performed according to the manufacturer’s instructions, using afinal concentration of 250 nM and 30 minutes of incubation at 37 C.

Microscopy was performed using an Olympus IX81 wide field microscope, orLeica TCS-SP2 or Zeiss LSM 510/FCS confocal microscopes. Images wereprocessed using NIH ImageJ version 1.42 (www.rsbweb.nih.gov/ij). Samples wereprepared and 3D-SIM (Microbial Imaging Facility, University of Technology,Sydney) performed as previously described (Yeoman et al., 2011). Analysis of thedimensions of at least seven cells from DIC images was performed using ImageJ.Isosurface renderings of GAP50–GFP-expressing gametocytes were generatedusing IMOD software (Kremer et al., 1996; Mastronarde, 1997).

Immunoprecipitation and solubility studies

Gametocytes were harvested on days 3, 5, 7, 9 and 11, representing stages I to V.Pellets of magnet-purified gametocytes were lysed in 0.015% saponin on ice for15 minutes. The saponin pellets were solubilized in SDS loading buffer and equalamounts (10 mg protein) were separated by SDS-PAGE and transferred tonitrocellulose for immunoblotting.

For solubility studies, magnet-purified stage IV gametocytes were lysed in0.015% saponin and the pellets incubated for 15 minutes with 1% Triton X-100or RIPA buffer (1% IGEPAL CA-630), 150 mM NaCl and 0.5% sodiumdeoxycholate on ice, or with 0.1% SDS and 50 mM TRIS, pH 7.4 at roomtemperature or with 2% SDS at room temperature. The soluble supernatant andinsoluble pellet fractions were collected. Equivalent amounts of protein wereseparated by SDS-PAGE and transferred to nitrocellulose for immunoblotting.

For immunoprecipitation studies, purified parasites were lysed in RIPA bufferfor 15 minutes on ice in the presence of protease inhibitor, centrifuged, and thesupernatants collected. The precipitating agents were GFP–TRAP reagent(Chromotek) and Protein A Sepharose and anti-rabbit GAP45 (1:500) (Yeomanet al., 2011). The pellets were analyzed by SDS-PAGE and immunoblottingblotting.

Immunoblotting was performed as previously described (Dixon et al., 2011).Briefly, all antibodies were prepared in PBS containing 3.5% skim milk, and bothprimary and secondary incubations were performed for 1 hour. Three washes ofPBS containing 0.1% Tween-20 were performed between each incubation. Thefollowing primary antibodies were used: mouse anti-GFP (1:500, Roche), rabbitanti-GFP (1:500), rabbit anti-GAP45 [1:500 (Baum et al., 2006b)], rabbit anti-GAP50 [1:500 (Baum et al., 2006b)], mouse anti-GAP50 [1:500 (Baum et al.,2006b)], mouse anti-Pfs16 [1:1000 (Dixon et al., 2009)], mouse anti-MTIP [1:500(Green et al., 2006)] and mouse mAb anti-AMA1 [mAb 1F9 1:500 (Coley et al.,2001)]. Goat anti-mouse or anti-rabbit IgG secondary antibodies conjugated tohorseradish peroxidase were used. Electrochemiluminescence reagents andautoradiography were used to visualize the immunoblots.

Cell deformabilityMature stage asexual (30–36 hour trophozoite) and stage III–V gametocytes weremagnet-purified from culture, washed in PBS, then reconstituted with uninfectedRBCs to obtain a working parasitemia of 70% (Maier et al., 2008). The cellsamples (5 ml) were mixed with 500 ml of polyvinylpyrrolidone (PVP) solution ata viscosity of 25 mPa second (Rheo Meditech). The elongation index wasmeasured in a RheoScan Ektocytometer according to the manufacturer’s

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instructions. Measurements were acquired over the 0–20 Pa range. Values shown

are means6s.e.m. from a minimum of three individual measurements. The results

shown are from a single experiment. The experiment was repeated showing

comparable results (two separate experiments).

Electron microscopy and tomography

Transmission electron microscopy and tomography of asexual stages were

performed as previously described (del Pilar Crespo et al., 2008; Hanssen et al.,

2010); for merozoites, the cells were post-fixed with potassium-ferricyanide-reduced osmium tetroxide. Gametocytes from selected stages were fixed and

embedded using a modification of a published protocol (Kass et al., 1971). Briefly,

cells were fixed in 5% glutaraldehyde in PBS, pH 7.3, overnight, then embedded

in 3% agarose before undergoing post-fixation of lipids in osmium tetroxide (1%

in PBS) for 1 hour followed by progressive dehydration in ethanol. Samples were

further dehydrated using progressive ethanol:acetone ratios of 2:1, 1:1 and 1:2, and

finally immersed in acetone for 10 minutes. Acetone was gradually replaced with

liquid epoxy resin, and samples embedded and stained as described above. Forboth asexual and gametocyte samples, 70-nm thick sections were observed at

120 keV using a JEOL 2010HC transmission electron microscope (EM Unit, La

Trobe University,). For electron tomography, sections of 200 or 300 nm were cut

and stained and observed at 200 keV on an FEI Tecnai G2 F30 electron

microscope (Bio21 Institute, Melbourne). Tilt series were collected every 2˚between 270˚ and +70˚ and reconstructed to generate cell models as described

previously (Abu Bakar et al., 2010; Yeoman et al., 2011). Tomograms were

generated using the IMOD software (Kremer et al., 1996; Mastronarde, 1997).

IMOD was also used to generate segmentation models where contours wereassigned manually.

Cryo-electron tomography

Gametocytes were transferred onto carbon-coated grids. After removal of excessliquid, grids were rapidly frozen by plunging into liquid ethane then transferring to

liquid nitrogen. Grids were mounted on a Gatan cryo-holder and imaged using a

Tecnai G2 F30 (300 keV). Tilt series were collected every 2˚ between 270˚ and

+70 . The total electron dose kept was 6000 e2/nm2.

AcknowledgementsWe would like to thank Alex Lowdin, Michael Jones and MichaelJohnson for assistance with microscopy protocols. We thankTony Holder (National Institute for Medical Research, London)for donating anti-MTIP antibodies, Robin Anders (La TrobeUniversity) for anti-AMA1 antibodies, Jake Baum (Walter andEliza Hall Institute) for the anti-GAP50, GAP45 and MyoA antiseraand David Baker (London School of Hygiene and TropicalMedicine) for anti-Pfs16 antibodies. We thank Tim Gilberger andTobias Spielmann (Bernhard Nocht Institute) for useful discussions.

FundingM.K.D. is supported by an Australian Postgraduate Award (APA).L.T. is an ARC Australian Professorial Fellow. M.W.A.D. issupported by an NHMRC training fellowship. C.B.W. is aNHMRC Senior Research Fellow. This work was supported bygrants from the ANZ Group of Trustees, the Ramaciotti Foundationand an ARC Discovery project.

Supplementary material available online at

http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.099002/-/DC1

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