Release of Hepatic Plasmodium yoelii Merozoites into the Pulmonary Microvasculature

Abstract

Plasmodium undergoes one round of multiplication in the liver prior to invading erythrocytes and initiating the symptomatic blood phase of the malaria infection. Productive hepatocyte infection by sporozoites leads to the generation of thousands of merozoites capable of erythrocyte invasion. Merozoites are released from infected hepatocytes as merosomes, packets of hundreds of parasites surrounded by host cell membrane. Intravital microscopy of green fluorescent protein-expressing P. yoelii parasites showed that the majority of merosomes exit the liver intact, adapt a relatively uniform size of 12-18 microm, and contain 100-200 merozoites. Merosomes survived the subsequent passage through the right heart undamaged and accumulated in the lungs. Merosomes were absent from blood harvested from the left ventricle and from tail vein blood, indicating that the lungs effectively cleared the blood from all large parasite aggregates. Accordingly, merosomes were not detectable in major organs such as brain, kidney, and spleen. The failure of annexin V to label merosomes collected from hepatic effluent indicates that phosphatidylserine is not exposed on the surface of the merosome membrane suggesting the infected hepatocyte did not undergo apoptosis prior to merosome release. Merosomal merozoites continued to express green fluorescent protein and did not incorporate propidium iodide or YO-PRO-1 indicating parasite viability and an intact merosome membrane. Evidence of merosomal merozoite infectivity was provided by hepatic effluent containing merosomes being significantly more infective than blood with an identical low-level parasitemia. Ex vivo analysis showed that merosomes eventually disintegrate inside pulmonary capillaries, thus liberating merozoites into the bloodstream. We conclude that merosome packaging protects hepatic merozoites from phagocytic attack by sinusoidal Kupffer cells, and that release into the lung microvasculature enhances the chance of successful erythrocyte invasion. We believe this previously unknown part of the plasmodial life cycle ensures an effective transition from the liver to the blood phase of the malaria infection.

Full text

Release of Hepatic Plasmodium yoelii
Merozoites into the Pulmonary
Microvasculature
Kerstin Baer
1
, Christian Klotz
1
, Stefan H. I. Kappe
2
, Thomas Schnieder
3
, Ute Frevert
1*
1 Department of Medical Parasitology, New York University School of Medicine, New York, New York, United States of America, 2 Seattle Biomedical Research Institute,
Seattle, Washington, United States of America, 3 Department of Parasitology, University of Veterinary Medicine, Hannover, Hannover, Germany
Plasmodium undergoes one round of multiplication in the liver prior to invading erythrocytes and initiating the
symptomatic blood phase of the malaria infection. Productive hepatocyte infection by sporozoites leads to the
generation of thousands of merozoites capable of erythrocyte invasion. Merozoites are released from infected
hepatocytes as merosomes, packets of hundreds of parasites surrounded by host cell membrane. Intravital microscopy
of green fluorescent protein–expressing P. yoelii parasites showed that the majority of merosomes exit the liver intact,
adapt a relatively uniform size of 12–18 lm, and contain 100–200 merozoites. Merosomes survived the subsequent
passage through the right heart undamaged and accumulated in the lungs. Merosomes were absent from blood
harvested from the left ventricle and from tail vein blood, indicating that the lungs effectively cleared the blood from
all large parasite aggregates. Accordingly, merosomes were not detectable in major organs such as brain, kidney, and
spleen. The failure of annexin V to label merosomes collected from hepatic effluent indicates that phosphatidylserine
is not exposed on the surface of the merosome membrane suggesting the infected hepatocyte did not undergo
apoptosis prior to merosome release. Merosomal merozoites continued to express green fluorescent protein and did
not incorporate propidium iodide or YO-PRO-1 indicating parasite viability and an intact merosome membrane.
Evidence of merosomal merozoite infectivity was provided by hepatic effluent containing merosomes being
significantly more infective than blood with an identical low-level parasitemia. Ex vivo analysis showed that
merosomes eventually disintegrate inside pulmonary capillaries, thus liberating merozoites into the bloodstream. We
conclude that merosome packaging protects hepatic merozoites from phagocytic attack by sinusoidal Kupffer cells,
and that release into the lung microvasculature enhances the chance of successful erythrocyte invasion. We believe
this previously unknown part of the plasmodial life cycle ensures an effective transition from the liver to the blood
phase of the malaria infection.
Citation: Baer K, Klotz C, Kappe SHI, Schnieder T, Frevert U (2007) Release of hepatic Plasmodium yoelii merozoites into the pulmonary microvasculature. PLoS Pathog 3(11 ):
e171. doi:10.1371/journal.ppat.0030171
Introduction
Two billion people, more than one third of the world’s
population, live at risk for malaria and about 1 billion are
infected. Each year there are 300 million to 500 million new
cases with 2–3 million deaths, the vast majority youn g
children in Africa. We are now forty years past the discovery
that radiation-attenuated sporozoites protect against malaria
[1], but we still lack an efficient malaria vaccine to combat this
deadly parasitic disease, and drug resistance is wide-spread
[2].
The malaria infection begins with the introduction of
sporozoites from the bite of an infected Anopheles mosquito
[3,4]. The sporozoites travel to the liver and develop in
hep atocytes to large exoerythrocytic forms (EEFs) [5,6].
Schizogonic division of the EEF then results in the formation
of thousands of first-generatio n merozoites, which are
responsible for the initiation of clinical malaria. Merozoites
have a short life span and must infect erythrocytes immedi-
ately after release into the bloodstream [7]. Merozoites are
also highly susceptible to phagocytosis and must therefore
avoid contact with macrophages [8]. Acute danger of
phagocytic elimination is presented in the form of Kupffer
cells [8], the resident phagocytes of the liver that comprise by
far the largest population of tissue macrophages of the body
[9]. Kupffer cells are predominantly located at sinusoidal
bifurcations, largely within and often spanning the sinusoidal
lumen [9–11], thereby presenting significant obstacles for
non-self particulate material. This strategic position of
Kupffer cells makes it difficult for free merozoites to exit
the liver without being trapped by these surveillance cells of
the innate immune system.
The first evidence suggesting that merozoites can be
released from hepatocytes as clusters, held together by host
cell cytoplasm, was presented several decades ago in Garn-
Editor: L. David Sibley, Washington University School of Medicine, United States of
America
Received June 20, 2007; Accepted September 26, 2007; Published November 9,
2007
Copyright: Ó 2007 Baer et al. This is an open-access article distributed under the
terms of the Creative Commons Attribution License, which permits unrestricted
use, distribution, and reproduc tion in any medium, provided the original author
and source are credited.
Abbreviations: ASGR1, asialoglycoprotein receptor 1; EEF, exoerythrocytic form;
GFP, green fluorescent protein; PI, propidium iodide; PS, phosphatidyl serine; PV,
parasitophorous vacuole; PVM, parasitophorous vacuole membrane; PyGFP, GFP-
expressing Plasmodium yoelii parasite
* To whom correspondence should be addressed. E-mail: ute.frevert@med.nyu.edu
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ham’s ultrastructural examination of Plasmodium yoelii–in-
fected murine livers and described in more detail in Meis’
extensive electron microscopic studies on P. berghei infection
of the mouse [5,12,13]. More recently, we and others reported
that merozoites are released as ‘‘ extrusomes’’ or ‘‘ mero-
somes’’ that contain hundreds to thousands of parasites
[14,15] (reviewed in [16,17]). Our initial intravital observa-
tions using green fluorescent P. yoelii and BALB/c mice
revealed extensive movement within EEFs nearing comple-
tion of merozoite maturation culminating in budding and
release of merosomes into the hepatic bloodstream [14]. An
elegant series of in vitro studies described the differentiation
of P. berghei merozoites in the human hepatoma cell line
HepG2 [15]. While developing into hepatic schizonts, the
intracellular parasites prevent the initiation of a death
program in their host cells, but leave them to die once
merozoite formation is comp lete. Underlying molecular
details remain to be determined, but the data suggest that
host cell death in this in vitro model shares more features
with autophagy than apoptosis or necrosis [18]. However,
information on the viability of hepatocytes releasing mer-
ozoites into the sinusoidal blood is lacking to date.
Because P. yoelii infection of the mouse represents an
accepted model closely reflecting human malaria [19], we
used a variety of microscopic techniques to s tudy the
dynamics of merosome budding from infected hepatocytes
and the fate of hepatic merozoites in the body. Confocal
images provided measurements of merosome volume, mer-
osomal merozoite content, and EEF volume, and appropriate
mathematic processing of these data allowed us to calculate
the number of hepatic merozoites produced by P. yoelii
sporozoites in the murine host. Using intravital and ex vivo
microscopy, we found that the vast majority of hepatic P. yoelii
merozoites leave the liver camouflaged as merosomes,
disseminate within the cardiovascular system, and arrest in
the lungs. Molecular markers revealed that merosomal
merozoites remain viable and infectious until being released
into the pulmonary microcirculation. In contrast, various in
vivo and ex vivo assays suggest that unreleased merozoites
and the exhausted host cell eventually succumb to necrosis.
The resulting inflammatory stimulus attracts neutrophils, and
mononuclear phagocytes thus give rise to the formation of
microgranulomata. Overall, this systematic temporal and
quantitative analysis indicates that merosome formation and
release by host hepatocytes, merosome transport to and
sequestration in the lungs, and release of merozoites into the
pulmonary microvasculature are parts of a previously
unrecognized phase of the Plasmodium life cycle.
Results
Morphology of Late P. yoelii Liver Stages
P. yoelii–infected mice have been suggested to represent a
suitable model for human malaria [20]. We also consider P.
yoelii an appropriate rodent model for liver stage analysis
because it induces less inflammation in murine livers than P.
berghei and produces more EEFs [21], which in addition are
generally larger and contain more meroz oites [12,22,23]
(Table S1). While available for other species such as P. berghei,
information is scarce regarding ultrastructural changes
during P. yoelii EEF maturation in the liver and the
subsequent release of first generation merozoites [12,24]. To
help fill this gap and to expand our previous investigation of
Plasmodium merosomes in live mice [14], we used several light
and electron microscopy techniques to examine this process.
Mature Plasmodium EEFs contained thousands of merozoites
enclosed in a parasitophorous vacuole (PV). Up to the final
developmental stage and onset of merozoite release, infected
hepatocytes remained in close contact with neighboring
uninfected parenchymal (Figure 1A and 1B) and sinusoidal
cells (Figure 1C). Shortly before merosome formation, the PV
membrane (PVM) disintegrated so that host cytoplasm
contained a mixture of mature merozoites, morphologically
intact hepatocyte organelles (Figure 1D), parasite remnant
bodies (or pseudocytomeres [5]), and parasite stroma left over
from schizogonic merozoite formation (Figure 1E). Some of
the sinusoids adjacent to infected hepatocytes remained filled
with erythrocytes indicating preservation of function, but
others were compressed by the expanding parasite and lacked
erythrocytes suggesting local obstruction of blood flow
(Figure 1C). To calculate the merozoite content of mature
EEFs (see below), we needed accurate measurements of the
EEF size. Compared to tissue sections, intravital microscopy
of green fluorescent protein (GFP) Plasmodium yoelii parasite
(PyGFP)–infected mouse livers (Figure 1F) offe red the
advantage of examining live tissue within an intact animal,
thus avoiding artifacts associated with both fresh and fixed
sections. Mature EEFs within the liver typically have a slightly
ellipsoid shape with the minimum and maximum diameters
ranging from 40 to 75 lm (with averages of 49.2 6 10.3 lmto
55.6 6 9.0 lm), respectively (n ¼ 16).
Mechanism of Exoerythrocytic Merozoite Release
Detailed intravital examination of 30 mice at times ranging
from 30 to 74 h after intravenous infection with PyGFP
sporozoites allowed us to follow the complex series of events
involved in merozoite liberation from hepatocytes. We
monitored more than 60 EEFs over this period and observed
the earliest merosome budding at 46 h (Figure 2A, Videos S1–
S3), a time in general agreement with earlier work reporting
the first appearance of P. yoelii in the blood at 45.5 h [12]. Of
these 60 EEFs, 20 reached maturity during the observation
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Plasmodium Merozoite Release in the Lung
Author Summary
The malaria parasite Plasmodium undergoes one large round of
multiplication in the liver before beginning the blood phase of the
life cycle, the phase that causes the typical episodes of fever and
chills. Using intravital microscopy and fluorescent parasites, we
studied the mode and dynamics of parasite release from the liver, a
critical stage in the malaria life cycle. Earlier work had indicated that
infected liver cells could release packets of dozens to hundreds of
parasites enveloped by host cell membrane, structures now known
as merosomes. We report here that this is the predominant
mechanism of parasite release from the liver. The host-derived
merosome membrane lacks a marker for phagocytic engulfment,
thus allowing safe passage through the gauntlet of Kupffer cells,
highly active liver macrophages. Merosomes remain intact during
passage through the heart and become sequestered within lung
capillaries where the membrane eventually disintegrates liberating
the parasites into the lung circulation. We propose that this
previously unknown part of the life cycle of Plasmodium facilitates
red blood cell invasion, thus jump-starting the blood phase of the
life cycle and the onset of clinical malaria.

period and released merozoites, while the rest remained
immature. The majority (13) of these 20 EEFs released
merozoites by merosome formation. Merosome formation
continued until 56 h after infection, thus confirming the
asynchronous nature of P. yoelii EEF maturation, a common
observation in Plasmodium-infected livers [5,25]. Because we
infected by intravenous sporozoite injection, the well-known
slow release of sporozoites from the mosquito bite site [3,26]
alone cannot account for the asynchronicity observed here.
For individual EEFs, the process of merosome budding and
release lasted several hours during which time the host cell
gradually decreased in size and separated from neighboring
cells (Figure 2B). In addition to fully formed green fluorescent
merozoites, released merosomes contained non-fluorescent
remnant bodies and host cell organelles, thus providing
further evidence that merosome budding occurs after
rupture of the PVM. Eventually, the host cell membrane
appeared to lose its integrity and allowed some leftover
merozoites to enter the bloodstream singly and without
protection by a merosomal membrane (Figure 2C and 2D,
Video S4). GFP radiated out from the disintegrating EEF into
the surrounding tissue, implying that parasite antigens and
host cell cytoplasm were set free as well. Indeed, electron
microscopic examination showed free mitochondria in the
sinusoidal lumen (Figure 2E). Size and shape of these
organelles revealed hepatocyte origin. Eventually, inflamma-
Figure 1. Characterization of Late P. yoelii EEFs
(A) Semithin Epon section showing a mature EEF containing thousands of merozoites at 52 h after P. yoelii sporozoite infection. Note that few sinusoids
are visible surrounding the EEF. Bar ¼ 10 lm.
(B) An infected hepatocyte containing a mixture of merozoites and host cell organelles indicating that the PVM has ruptured. The membrane of the
infected cell is in close contact (arrows) with a neighboring hepatocyte. H, hepatocyte. *, merozoites. Bar ¼ 5 lm.
(C) Merozoites lie in close apposition to ultrastructurally well-preserved host cell mitochondria. M, mitochondria; *, merozoites. Bar ¼ 1 lm.
(D) EEF containing numerous merozoites and a few host cell organelles and remnant bodies. Note that one adjacent sinusoid has collapsed (arrows),
while another contains an erythrocyte in its lumen indicating preserved blood flow. RB, remnant bodies. Bar ¼ 5 lm.
(E) Merozoites and remnant bodies of various shapes and sizes are embedded in a loose matrix. RB, remnant bodies; *, merozoites. Bar ¼ 5 lm.
(F) Intravital micrograph showing a mature PyGFP EEF at the onset of merosome formation. The nuclei of the host cell and the neighboring hepatocytes
are visualized by Hoechst staining. Bar ¼ 10 lm.
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Plasmodium Merozoite Release in the Lung

tory cells were attracted to the site of the disintegrating EEF.
During phagocytic removal of debris from dead merozoites
and host cells, neutrophil granulocytes and mononuclear
phagocytes transformed the site of the former EEF into a
small granuloma (Figure S1), a structure commonly reported
at late stages of Plasmodium liver infection [5,8,27–31]. Thus,
merosome formation in the liver occurs over a period of
about 10 h and is followed by disintegration of the host cell
and some leftover parasites, clearance of the remains by
infiltrating phagocytes, and production of a small granuloma.
Merozoites were also liberated by a less frequent mecha-
nism. Starting earlier than merosome formation (42 h post
inoculation), some infected hepatocytes rapidly discharged
their content of merozoites and cell organelles by a
Figure 2. Merozoite Release by Merosome Formation
(A) Intravital microscopy of merosomes budding from an EEF. Note that a few individual merozoites are located in the vicinity of the EEF (see Videos S1–
S3). Bar ¼ 10 lm.
(B) Semithin Epon section showing an EEF that has lost contact with the surrounding parenchyma (arrows). Bar ¼ 10 lm.
(C) Projection of an EEF in the process of disintegration; note that the cell releases individual merozoites into the environment (see Video S4). Bar ¼ 10
lm.
(D) Electron micrograph showing merozoites located free in a liver sinusoid. The apical pole of one of the merozoites is in close contact with an
erythrocyte (insert). E, erythrocyte; *, merozoites. Bar ¼ 1 lm.
(E) Liver sinusoid containing free hepatocyte mitochondria (arrows) and a free merozoite. E, erythrocyte; *, merozoite. Bar ¼ 5 lm.
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Plasmodium Merozoite Release in the Lung

mechanism appearing to involve rupture of the cell mem-
brane (Figure 3A–3E, Video S5). In some cases, the process
was complete in as little as 5 min; in others it lasted as long as
60 min. Of the EEFs rupturing in this manner, 80% harbored
mature merozoites, but 20% had a homogeneous cytoplasm;
thus, schizogony had not even begun (Figure S2). Occasion-
ally, electron micrographs show ed immature merozoites
incompletely separated from remnant bodies yet released
into the sinusoidal bloodstream (Figure 3F). This apparent
rupture-release left large faintly fluorescent EEF ghosts at the
site of the former host cell. Because our intravital observa-
tions were based on confocal microscopy, we considered the
possibility of phototoxicity playing a role in this rupture-
release mechanism. However, since EEF ghosts identical to
those resulting from observed rupture were detectable at the
very beginning of intravital examination, we could reject that
possibility. Because the EEFs did not decrease in volume prior
to transformation into a ghost, and we did not find
erythrocytes associated with these ghosts, we suspect that
the remains of the host cell cytoskeleton, the surrounding
extracellular matrix, and/or the sinusoidal cell layer resealed
the ghost after merozoite release; thus, preventing the
formation of hemorrhages. Similar to the end of the
merosome release mechanism (see above), EEF ghosts were
infiltrated by inflammatory cells that gave rise to small
granulomata. When we combine results from intravital
microscopy, showing that both mature and immature EEFs
undergo this rapid decay, with our electron microscopy data,
showing that some of the rupturing EEFs were immature, we
conclude that this rapid release process is a result of abortive
EEF development that, in the absence of host cell membrane
protection, exposes the parasites to Kupffer cell phagocytosis.
To demonstrate that merozoites within merosomes are
alive and to help exclude the possibility that merosome
release represents an abnormal development, we injected
infected mice with markers that reveal cell viability in vivo. At
points ranging from 51 to 74 h post inoculation, mice were
injected with a mix of the membrane-permeable DNA stain
Hoechst 33342 and the dead cell marker propidium iodide
(PI). Subsequent intravital confocal microscopy revealed that
PI does not enter merosomes or intact EEFs (Figure 4A and
4B), but does stain some of the merozoites left behind in EEF
ghosts and also in EEFs that had disintegrated after
merosome budding (Figure 4C and 4D). These findings
support the interpretations above in that they suggest that
merozoites that fail to exit the host cell eventually succumb to
necrosis.
Efforts to determine the mode by which merosomes breach
the sinusoidal cell layer failed so far due to insufficient
numbers of suitable events for analysis. We suspect that
budding occurs through the endothelial fenestration rather
by a paracellular route, because of the extreme natural
variability of the diameter of the fenestrae in response to
changes in blood pressure and other physiologic stimuli.
Interestingly, mature EEFs were frequently surrounded by a
layer of flattened cells that had incorporated the dead cell
stain PI (Figure 4A and 4B). Perhaps the death of these cells is
due to extreme compression by the extensive expansion of
the EEF during the final stage of development. Occasionally,
merosomes were found budding into such dead cells, but the
Figure 3. Rapid EEF Decay
(A–D) Individual frames from an intravital video showing the process of liver stage transformation into an EEF ghost (see Video S5). (A) The still image
shows an intact EEF adjacent to a ghost. G, ghost.
(B) Individual frame showing rapid decay of the EEF shown on the left of (A). Note that loss of cytosolic GFP reveals the presence and arrangement of
the merozoites (arrow).
(C–D) Only a few of the thousands of merozoites remain visible in the faintly fluorescent EEF ghost.
(E) Eventually, the infected hepatocyte has transformed into an EEF ghost. Note that both ghosts retain close contact with the surrounding tissue (A–E).
Bars ¼ 20 lm.
(F) Electron micrograph showing a liver sinusoid with merozoites that are incompletely separated from a remnant body. RB, remnant body; G, ghost; *,
merozoite. Bar ¼ 1 lm.
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immobility of the parasites indicated that they were trapped
(Figure 4A).
Dynamics of Merosome Budding and Transport
When hepatic merosomes initially bud from infected
hepatocytes (Figure 5), they are highly variable in size and
contain hundreds to thousands of mature merozoites, while
merosomes in blood draining from the liver were smaller and
more uniform in size. Intravital microscopy showed very large
merosomes moving far more slowly than small ones, which
leave the liver lobules at a velocity close to that of blood cells
(unpublished data). We frequently observed meros omes
hindering the free flow of the blood as they moved along a
sinusoid (Figure 5C–5G) as well as being hindered by the
vascular architecture. The speed of merosome transport at
any instant depended on the diameter and local structure of
the sinusoid as well as the size of the merosome. We recorded
large merosomes being arrested at sinusoidal bifurcations
where they occasionally even reversed direction of movement
(Figure 5A and 5B and Videos S6 and S7). Because
morphological measurements taken in vitro are subject to
artifact and do not reveal in vivo dynamics, we sought a better
understanding of sinusoidal architecture using intravital
analysis of uninfected transgenic Tie2-GFP mice that have
fluorescent vascular endothelia [32]. We found sinusoidal
diameters to range from 3.4 lm to 14.1 lm (6.7 6 1.9 lm; n ¼
94) under normal blood pressure conditions. Although large
merosomes greatly exceed this size range, their considerable
deformability allowed them to gradually wind their way
towards the central vein and exit the liver without rupture
and release of merozoites, a process aided by resizing (Figure
5A and 5B). We occasionally observed large merosomes
subdividing into smaller ones while traveling through sinus-
oids (Videos S8 and S9), but we suspect that shear forces
associated with the faster blood velocity in larger vessels
caused merosomes in the hepatic effluent and inside lung
Figure 4. Hepatocyte and Merozoite Viability during Merosome Formation
At 52–70 h after infection with PyGFP, mice were intravenously injected with a mixture of the membrane-permeable DNA stain Hoechst 33342 and the
dead cell dye PI prior to intravital microscopic examination. (A) Confocal scan showing an intact EEF that has not incorporated PI (red) and is therefore
considered viable. Note that merozoites have been discharged into surrounding cells (arrow) some of which have taken up the red dead cell stain.
(B) Due to merosome formation, this EEF has decreased in size and detached from the surrounding tissue. The neighboring cells appear to be
compressed and are stained with PI (red).
(C) A few merozoites are left behind in an EEF that has disintegrated after repeated merosome budding. The two nuclei of the host cell (arrows) have
incorporated predominantly PI (red) and some Hoechst stain (blue) and appear pink. In addition, some of the merozoites have lost their green
fluorescence and appear red due to PI staining, while others have retained GFP and excluded PI and are therefore viable. The nuclei of the neighboring
hepatocytes are visualized by Hoechst staining (blue).
(D) EEF at a late stage of disintegration. Some remaining merozoites are viable (green), while other merozoites are dead (red). The nuclei of some
surrounding cells have also incorporated PI (arrows). Bars ¼ 10 lm.
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capillarie s to be generally small er and uniform in size
compared to those in the liver. The importance of mechan-
ical forces for resizing is demonstrated by another set of
experiments in which PyGFP-infected mouse livers were
removed from the animals and analyzed ex vivo by confocal
microscopy, i.e., in the absence of blood flow. The sinusoids of
such livers contained merosomes of an unusually large size
(Figure S3A). When livers were perfused with medium prior
to ex vivo confocal microscopy, the sinusoids contained even
larger merosomes (Figure S3B). We contend that lack of
blood flow prevents subdivision of large merosomes into
smaller ones and that liver perfusion hastened merosome
budding and liberation from the host cell.
Merosome formation results in packaging a mixture of
parasites, remnant bodies, and host cell cytoplasm within host
cell membrane for release into the sinusoidal lumen (Figures
5H and 6A). Ultrastructurally, the merosomal matrix con-
tained well-preserved merozoites and morphologically intact
host cell mitochondria (Figure 6B) suggesting that these
organelles are viable at the time of merosome budding.
Merosomes also typically contained remnant bodies (Figure
6A) suggesting that these leftovers from EEF schizogony
represent a natural component of the merosomal cytoplasm.
In the absence of better viability markers, we interpret the
presence of MSP-1 on the surface of merozoites in both
mature EEFs and merosomes (Figure 6D and 6E) to indicate
intactness and complete differentiation of the parasites, and
propose that merosomes are linked to productive infection of
erythrocytes. Disintegration of the PVM prior to merosome
formation indicates the merosome membrane is derived from
hepatocyte cell membrane. Asialoglycoprotein receptor 1
(ASGR1), a protein expressed only on parenchymal liver cells
[33–37], was detectable by immunofluorescence lining the
basal hepatocyte surface within the space of Disse (Figure 6C).
ASGR1 clearly surrounded mature EEFs (Figure 6D), but it
was absent from the merosomal membrane (Figure 6E). The
Figure 5. Dynamics of Merosome Transport in the Liver
(A) Multiple merosomes separate from an EEF and are rapidly transported towards the central vein (see Video S6). Bar ¼ 10lm.
(B) Large merosomes move slowly within the sinusoidal lumen (see Video S7). Remnant bodies can be differentiated from merozoites by their larger size
and lack of GFP (arrows). Bar ¼ 10 lm.
(C–G) Individual frames of an intravital confocal video showing two smaller merosomes gliding along a sinusoid. Bars ¼ 10 lm.
(H) Semithin liver section 52 h after infection with P. yoelii. Multiple merosomes, presumably released from a single EEF, can be found inside the
sinusoids (arrows). Note the presence of remnant bodies in merosomes. Bar ¼ 10 lm.
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Plasmodium Merozoite Release in the Lung

lack of this hepatocyte surface protein could be due to
dedifferentiation of the infected host cell or modification by
the intracellular parasite at late stages of EEF maturation.
However, because the ASGR1 label was located predom-
inantly in the space of Disse rather than on the hepatocyte
membrane, more work is needed to define the composition of
the merosome membrane.
Merosomes Exit the Liver Intact
Because intravital observations showed that merosomes
remain intact during transport towards the central vein, we
examined the hepatic venous effluent for membrane-envel-
oped parasites. To do this, we opened the inferior Vena cava
at its point of entry into the diaphragm and collected blood
from the peritoneal cavity. Thick smears were prepared from
5 ll blood, and the concentrations of venous merosomes from
three separate experiments were measured. In hepatic venous
blood collected 52 h post infection with 2.5 3 10
6
sporozoites,
we found 28.7 6 4.3 merosomes per ll, and 69% of these
merosomes contained between 100 and 200 merozoites
(unpublished data).
While much information is available on P. berghei–infected
HepG2 cells [15], in vivo data on the molecular composition
of the merosome membrane, for example phosphatidylserine
(PS) exposure, are lacking to date. To obtain more detail on
merosome structure, another set of experiments was per-
formed in which the parasite material available for examina-
tion was enhanced by liver perfusion. Beginning 52 h after
infection with PyGFP or wild-type (wt) P. yoelii, livers were
perfused with culture medium and the perfusate collected.
Cells were immobilized by attachment to Alcian blue–treated
glass-bottom dishes and immediately examined by confocal
microscopy using conditions that maintain viability. Perfu-
sate merosomes typically adapted a spherical shape in vitro
(Figure 7A) and 3-D images from confocal stacks demon-
strated a relatively uniform size containing several hundred
merozoites. Labeling by the phospholipid marker FM 4–64 FX
verified that the parasites were held together by a membrane
(Figure 7B).
Immediately after harvesting, the majority of the parasites
appeared viable and merosome membranes were negative for
annexin V labeling (Figure 7C); thus they do not display PS
that targets cells for phagocytosis. However, with increased
time in vitro, the presence of PS gradually became apparent
(Figure 7D). Merozoites in freshly isolated merosomes did not
stain with the dead cell marker PI, but those that became
positive for PS also lost the ability to exclude PI (Figure 7D). A
further viability assessment utilized YO-PRO-1, a DNA stain
that selectively passes through the (intact) plasma membrane
of apoptotic cells. Again, merozoites in freshly isolated
merosomes did not label with YO-PRO-1, but as time in
vitro progressed, they began to incorporate YO-PRO-1 along
with PI (Figure 7E and 7F). Within roughly 60 min of in vitro
examination, all merosomes were positive for annexin V, YO-
PRO-1, and PI. Attempts to quantify the time course of these
processes more precisely were prevented by the sensitivity of
the merosomes to the various steps of isolation from the liver
Figure 6. Merosomes Contain Viable Merozoites and Host Cell Organelles
(A) Semithin section showing an EEF releasing a merosome into a sinusoid. Note that remnant bodies are a normal component of the merosomal
cytoplasm. RB, remnant bodies. Bar ¼ 10 lm.
(B) Electron microscopy documents that merosomes contain well-preserved merozoites and hepatocyte mitochondria. E, erythrocyte; M, hepatocyte
mitochondria; *, merozoite. Bar ¼ 1 lm.
(C) Immunolabeling of frozen liver sections for ASGR-1 reveals the presence of the receptor on the basolateral portion of the membrane of all
hepatocytes (red). Bar ¼ 20 lm. Neither EEFs (D) nor merosomes (E, arrows) express ASGR-1 on their surface. The merozoites were visualized with
Hoechst (blue) and an antibody against MSP-1 in combination with PA-FITC (green). Bars ¼ 10 lm.
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Plasmodium Merozoite Release in the Lung

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Plasmodium Merozoite Release in the Lung

and concentration by centrifugation. Taken together, these
results suggest that P. yoelii merosomes leaving the liver
contain viable merozoites, and, similar to P. berghei–infected
HepG2 cells [15], lack PS as a membrane marker that signals
‘‘ eat-me’’ to phagocytes. Considering that Kupffer cells are
located largely within and often spanning the sinusoidal
lumen, thus presenting a significant obstacle for non-self
particulate material and damaged host cells [9–11], the lack of
PS on the merosome membrane is likely critical for merozoite
escape from this defense mechanism of the host.
Merosomes Accumulate in the Lung and Release
Merozoites
Because the entire hepatic effluent must pass through the
right ventricle and the pulmonary microcirculation before
reaching any other capillary bed, we suspected merosomes
might sequester in the lungs. To address this, we used ex vivo
confocal microscopy to examine the alveolar microvascula-
ture immediately after lung removal while the tissues were
intact and the cells alive. At time points from 46 to 58 h after
inoculation with PyGFP sporozoites, we found numerous
intact merosomes as well as individual parasites (Figure 8A–
8C). We did not find pulmonary merosomes earlier than 46 h
post inoculation nor later than 65 h, timing consistent with
our observation that merosome release begins and ends at
roughly 46 h and 56 h. MSP-1 labeling confirmed the maturity
of merozoites, both those within merosomes and those
already released into pulmonary capillaries (Figure 8D–8G
and Video S10). The small liver stage protein UIS-4, which
localizes to the PVM [38], was not detected (unpublished
data); thus providing more evidence that the PVM is not
involved in merosome formation [5]. As for merosomes in the
liver, pulmonary merosomes were negative by immunofluor-
escence for the hepatocyte receptor ASGR1 (Figure 8F and
8G).
In confocal images we often observed an asymmetric
arrangement of individual merozoites in relation to lung
merosomes and the pattern suggested some of the merosomes
were in the process of disintegrating and releasing merozoites
into the pulmonary microvasculature just as blood circu-
lation was stopped by lung removal (Figure 8B). Electron
microscopy supports the notion of merozoite release by
merosomal membrane degradation. Pulmonary merosomes
typically contained morphologically well-preserved mero-
zoites, but the cytoplasmic matrix was swollen, and host cell
organelles were clearly degenerating. The membrane of lung
merosomes was frequently disrupted or barely detectable
(Figure 8H) suggesting that free merozoites found in nearby
pulmonary microvasculature had just been released before
fixation (Figure 8I). The presence of erythrocytes containing
newly invaded merozoites (Figure 8J) supports the notion that
blood infection occurred in the lungs.
Merosomes Do Not Disseminate beyond the Lungs
To determine whether merosomes leaving the liver can
pass through the lungs and disseminate throughout the body,
we analyzed thick smears of blood collected from the aorta
and tail vein for merosomes. We also used intravital micro-
scopy, ex vivo imaging, and immunofluorescence microscopy
to examine capillary beds of spleen, kidney, and brain of the
same mice. While individual small parasites were occasionally
detectable, merosomes were completely absent from aorta
and tail vein blood and the microcirculation of these organs
(unpublish ed data). These results demonstrate effective
retention of hepatic merosomes in the lungs.
Merosomal Merozoites Are Viable and Infectious
At 52 h after infection with PyGFP, mice were injected with
Hoechst 33342 and PI, and the lungs were removed and
analyzed ex vivo. Confocal microscopy revealed that pulmo-
nary merosomes and free merozoites excluded PI (Figure 9)
and were also TUNEL-negative (unpublished data); thus
providing evidence of viability. Because infectivity is the
ultimate criterion for viability, we tested merosomal mer-
ozoites for their ability to induce a parasitemia in naı
¨
ve mice.
However, interpretation of results from inoculation with
blood containing merosomes is complicated by the presence
of infected erythrocytes. To circumvent this, we initially
attempted to eliminate parasitized erythrocytes using selec-
tive hypotonic lysis, but this also affected the integrity of the
merosomes. Our solution was to control for infected
erythrocytes by comparing the infectivity of two types of
blood taken from the same mouse: hepatic effluent (with
merosomes and some infected erythrocytes) and tail vein
blood (without merosomes, but with the same number of
infected erythrocytes). At 52 h after intravenous infection
with 2.5 3 10
6
wt P. yoelii sporozoites, hepatic effluent and tail
vein blood samples were collected for inoculation into other
mice. Parasitemia and merosome concentration were deter-
mined by analysis of thin and thick blood smears, respec-
tively. Preliminary studies showed that the parasitemia in
recipient mice injected with hepatic effluent blood rose
significantly faster compared to control mice injected with
tail vein blood (unpublished data) suggesting that merosomes
exiting the liver are infectious. This conclusion can be
confirmed once a method for merosome purification is
available.
Quantitative Analysis of Extrahepatic Merosomes
Because of the large number of parasites and their high
packing density, counting merozoites in EEFs is not feasible,
Figure 7. In Vitro Characterization of Hepatic Merosomes
(A) Merozoite nuclei in a PyGFP merosome were visualized with the membrane-permeable DNA stain SYTO-64 (red). Differential interference contrast
was used to capture the transmission image on the right of the panel.
(B) The phospholipid marker FM 4–64FX was used to visualize the membrane surrounding the merosome (red). Merozoite nuclei were stained with
Hoechst (blue).
(C) Immediately after harvesting from the hepatic effluent, the majority of merosomes were negative for annexin V (green) and PI (red).
(D) Prolonged in vitro cultivation led to PS exposure (green) in the outer leaflet of the merosomal membrane and to gradual incorporation of PI (red)
into individual merozoite nuclei. Parasite nuclei were visualized with Hoechst (blue).
(E and F) Freshly harvested merosomes excluded YO-PRO-1 (green), a DNA stain that selectively passes across apoptotic membranes, and the dead cell
stain PI (red). In vitro incubation of the merosomes led to the simultaneous uptake of both nucleic acid stains suggesting that merozoite death occurred
by necrosis rather than programmed cell death. Note that colocalization of YO-PRO-1, PI, and Hoechst causes nuclei to appear white. Bars ¼ 5 lm.
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Plasmodium Merozoite Release in the Lung

although estimates have been published [6,22,23]. We there-
fore used measurements obtained from merosomes to
calculate the number of merozoites contained in mature P.
yoelii EEFs. To do this, we first quantified the merozoite
content of a subset of smaller merosomes. We found that
merosomes with a diameter of 13.4 6 2.0 lm contained 134.7
6 51.6 merozoites. Then, we determined the average effective
volume merozoites take up inside merosomes. P. berghei
merozoites measure 1.0–1.2 3 1.5–1.7 lm [39–41], but because
the parasites are embedded in cytoplasm that also contains
parasite remnant bodies and host cell organelles, the effective
volume the parasites occupy is larger than their actual
volume of 0.78–1.23 lm
3
(based on the ellipsoid volume v ¼ 4/
3 p r
1
r
2
r
3
). Using a mathematical algorithm for optimal
packing of small spheres in a large sphere [42], we found that
P. yoelii merozoites have an effective diameter of roughly 2.2
lm and occupy an effective volume of 5.56 lm
3
in
merosomes. Because intravital and electron mi croscopy
showed that the merozoite packing density and the compo-
sition of the cytoplasm was basically identical in merosomes
compared to mature EEFs (after rupture of the PVM and
mixing of parasites and host cell organelles), we then used the
sphere packing algorithm to determine the merozoite
content of mature P. yoelii EEFs. Based on the measured
EEF diameter of 40–75 lm(seeabove),theeffective
merozoite diameter of 2.2 lm, and assuming a round EEF
Figure 8. Merosomes Accumulate in the Lungs and Release Merozoites into the Pulmonary Microcirculation
(A) Ex vivo confocal microscopy of a mouse lung 52 h after infection with PyGFP shows a large merosome located inside an alveolar capillary.
(B) A group of individual merozoites inside pulmonary capillaries.
(C) Small parasite aggregates and individual merozoites (arrowheads) fanning out in one direction from a merosome (arrow). This asymmetric parasite
distribution suggests that merosomes release merozoites into the pulmonary microcirculation. (A–C) Bars ¼ 10 lm.
(D–G) Mouse lung fixed 52 h after infection with P. yoelii. The frozen section was immunolabeled with MSP-1 followed by protein A-FITC (D, green)
indicating that the merozoites are intact. Labeling with goat-anti-ASGR-1 followed by rabbit anti-goat IgG-TX (E, red) revealed the absence of the
receptor on the merosomal membrane. Nuclei were visualized by Hoechst staining (F, blue).
(H) Electron micrograph showing well-preserved merozoites () in the lumen of an alveolar capillary. Only fragments of the merosomal membrane are
detectable (arrows). *, merozoite; A, alveolar lumen. Bar ¼ 1 lm.
(I) Free merozoites released into the lumen of a pulmonary capillary. *, merozoite; E, erythrocyte. Bar ¼ 1 lm.
(J) Alveolar capillary showing an erythrocyte (arrow) harboring a recently invaded merozoite in addition to several uninfected erythrocytes. *, merozoite;
E, erythrocyte. Bar ¼ 1 lm.
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Plasmodium Merozoite Release in the Lung

shape, we calculated that P. yoelii sporozoites produce 4,200–
29,000 hepatic merozoites (Table S1).
Discussion
We present here a new model for the transition from the
liver to the blood phase of the malaria life cycle (Figure 10):
large merosomes of various sizes bud from infected hepato-
cytes, enter the hepatic circulation, exit the liver intact,
subdivide into smaller more uniform sizes, but otherwise
withstand bloodstream shear forces during passage through
the right ventricle, and accumulate in the lungs where the
merosomes disintegrate and release merozoites to initiate the
erythrocytic phase of the malaria cycle. While EEF of avian
and reptilian malaria parasites develop in the reticulo-
endothelial or hematopoietic systems [43–45], a major evolu-
tionary change occurred with the mammalian malaria para-
sites, whose EEF mature in hepatocytes. Perhaps the
nutritionally rich and immunologically privileged hepatic
environment offers advantages, but it also presents a problem
for merozoites released from EEFs into hepatic sinusoids:
unless they invade an erythrocyte very quickly they face a
gauntlet of highly phagocytic Kupffer cells. The location of
most EEFs in the periportal area of the liver lobule [46] means
they must travel almost the full length of the sinusoid and
pass by a large complement of Kupffer cells before escaping
into relative safety outside the liver. As proposed previously
by us and others [14,15], our premise is that evolution
produced a countermeasure to this threat: release of
merozoites within large packets that are initially hidden from
the host’s innate immune system by envelopment with a
hepatocyte-derived membrane. Here we show that mero-
somes are delivered to the pulmonary microcirculation where
they are released. We propose that release of merozoites into
the lung microvasculature rather than into larger blood
vessels is advantageous, because the low macrophage density
and the reduced blood velocity with reduced shear forces will
enhance the ability of merozoites to invade erythrocytes.
Merosome disintegration in the lungs appears to be the
predominant mechanism of merozoite liberation into the
bloodstream for the following reasons: (1) In confirmation of
previous reports on the asynchronous nature of EEF
maturation [5,25], we observed P. yoelii merosome formation
in the liver from 46 h to 56 h after sporozoite infection.
Assuming a 10-h window of merosome release, roughly 3 ml
total blood volume in a 40 g mouse, and a 100% rate of
sporozoite infection and EEF development, 2.5 million
sporozoites would generate 4,167 maturing EEFs per minute,
corresponding to 1.4 merosome-releasing EEFs per ll blood.
(2) Assuming that extrahepa tic merosomes contain on
average 150 merozoites, the roughly 29 merosomes we found
per ll venous liver blood should have contained 4,350
merozoites. Since P. yoelii EEFs contain 4,200–29,000 mer-
ozoites (Table S1), up to 74% of the total number of
merozoites released by 1.4 EEFs per min and ll would have
been enclosed in merosomes. (3) A large number of
merosomes was arrested in alveolar capillaries suggesting
that many merosomes withstand the shear forces inside the
central cardiovascular system. Together, these data indicate
that a major proportion of the merosome population arrives
intact in the lungs and then gradually disintegrates, thus
liberating merozoites into the microvasculature. Pulmonary
merosomes were detectable in the lungs at least up to 58 h
after infection, i.e., beyond the period of release from the
liver (46–56 h), suggesting that they remained intact for at
least many minutes. Similar to hepatic merosomes, which
appeared to be infectious and did not stain with annexin V,
YO-PRO-1, or PI, pulmonary merozoites were ultrastructur-
ally well preserved, TUNEL-negative, and did not incorporate
PI. Together, these data suggest that merosomal merozoites
remain viable until their release into the pulmonary micro-
vasculature. Based on the above assumptions, we propose that
merozoite liberation in the lungs represents an integral part
of the Plasmodium life cycle.
Further support for our premise was found in the following
observations and suggestions derived from them. The notion
that merozoites shuttled out of the liver within merosomes
that are protected from phagocytosis by Kupffer cells [8] was
confirmed by demonstrating that murine Kupffer cells do not
phagocytoze PyGFP merosomes in vitro (unpublished data),
in agreement with the finding that P. berghei merosomes are
not ingested by a murine macrophage cell line in vitro [15].
Trager and Jensen’s finding that P. falciparum merozoite
invasion is enhanced by lack of flow and dense erythrocyte
packing [47,48] supports our hypothesis that merozoites
released within capillary beds have a better chance to invade
erythrocytes than those released into larger vessels. We can
imagine that capillary occlusion by arrested merosomes could
be helpful by causing local stagnation of the pulmonary blood
flow. We can also speculate that merosome arrest in lung
septal capillaries allows Plasmodium to exploit the unique
microenvironment of the blood-air barrier. Virtually nothing
is known about the biology of the first-generation (hepatic)
merozoites, but perhaps transient residence in the lungs
Figure 9. Pulmonary Merosomes Are Viable
Mouse lung analyzed ex vivo 48 h after infection with PyGFP. Generally,
neither merosomes nor free merozoites incorporated intravenously
injected PI indicating that the parasites were viable. Note that one of the
merozoites is PI-positive (arrowhead). Nuclei were visualized by Hoechst
staining (blue). Bar ¼ 10 lm.
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Plasmodium Merozoite Release in the Lung

provides these parasites with time and a suitable micro-
environment to gain infectivity for erythrocytes. The well-
oxygenated milieu of the terminal airways and the anasto-
mozed nature of the pulmonary microvasculature [49] likely
allow local occlusion of septal capillaries by merosomes
without causing the necrotic tissue damage associated with
infarction of microvessels in other organs.
Many aspects of the process of merosome formation and
release we describe are in agreement with earlier work, but
others are not. For example, we found that similar to P.
berghei–infected HepG2 cells, which detach in toto from the
culture vessel after merozoite differentiation is complete
[15], meros omes exiting P. yoelii–infected mouse livers
contain viable merozoites and initially do not expose PS
on their surface. This confirms earlier predictions [14,15]
that merozoites are safely shuttled out of the liver disguised
as merosomes. The presence of intact mitochondria in
mature EEFs indicates that Plasmodium liver stages are able
to manipulate hepatocytes in a way that useful organelles
(such as mitochondria as a source of energy) are preserved,
even after merosome budding. Our interpretation, namely
that Plasmodium controls certain host cell functions to the
Figure 10. Model of Merosome Dissemination and Merozoite Liberation
(A) After malaria sporozoite entry into a hepatocyte, the parasite begins to grow inside a PV to a size larger than its original host cell. Schizogonic
division results in the formation of thousands of erythrocyte-infective merozoites. During the final stage of differentiation, the PVM dissolves and allows
the parasites to mix with the remaining host cell organelles. Eventually, the plasma membrane of the infected hepatocyte bulges out and forms
merosomes; thus releasing merozoites, remnant bodies, and host cell mitochondria into the sinusoidal lumen. Camouflaged by host cell membrane,
merosomes are not recognized by Kupffer cells and are shuttled out of the liver. Infiltration of the remains of the infected host cell by mononuclear
phagocytes and neutrophil granulocytes gives rise to the formation of a small granuloma. Me, merosomes; RB, remnant bodies; Mi, host cell
mitochondria; KC, Kupffer cells; MB, mononuclear phagocytes; S, sporozoite.
(B) Shear forces inside the hepatic and other larger veins cause the merosomes to break down into smaller units. After passing through the right heart,
these small merosomes are arrested inside lung capillaries where (1) they eventually release infectious merozoites into the pulmonary microvasculature;
(2) local stagnation of the blood flow due to capillary occlusion by merosomes may result in dense erythrocyte packing; and thus (3) facilitate merozoite
infection. , alveolar space; H, hepatocyte; N, nucleus; E, endothelium; P-I, type I pneumocyte; P-II, type II pneumocyte; F, fibroblast; aMB, alveolar
macrophage; PMN, polymorphnuclear macrophage.
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Plasmodium Merozoite Release in the Lung

last minute, differs from the P. berghei HepG2 cell model, in
which the parasites induce death and detachment of their
host cells followed by merosome budding [15]. Further, the
cell membrane of P. yoelii–infected hepatocytes remains in
close apposition to that of neighboring parenchymal and
endothelial cells until the very end of EEF differentiation,
i.e., up to the onset of merosome budding, as reported
[5,12,13,50,51]. As merosomes are produced, the host cell
gradually decreases in size and loses contact with neighbor-
ing cells as reported [15]. We observed that after releasing
merosomes over several hours, the exhausted host cell
eventually disintegrates. Some free merozoites still escaped
and entered the sinusoidal lumen, thus being exposed to
attack by Kupffer cells. In contrast, others proposed that the
remaining host cell remnant is rapidly expelled in toto from
the tissue with the resulting void immediately filled by
neighboring cells [15,18]. We find that the necrotic remnant
attracts neutrophils and mononuclear phagocytes, which
eventually produce a small granuloma. Such granulomata
are a frequent observation in P. yoelii– and particularly in P.
berghei–infected mouse livers [5,8,27–31]. Rather than the
void created by expulsion of an EEF being filled quickly, our
in vivo observations suggest that hours, if not days, are
required for phagocytic removal of parasite and host cell
debris with subsequent repair of the structural damage
before normal tissue architecture is restored.
Although we found merosome formation to be the
predominant mode of merozoite release from the liver, we
observed a less frequent but still common alternative: EEFs
undergoing what we interpret as decay. This alternative
process of EEF ghost formation was rapid and typically
complete within minutes to an hour. In contrast to merozoite
release by merosome formation, ghost-forming EEFs did not
detach from the surrounding tissue. EEF decay was accom-
panied by leakage of GFP into the surrounding tissue
suggesting damage to the host cell membrane. It occurred
in immature EEFs (recognizable by a homogeneous green
fluorescent cytoplasm) and also in mature EEFs (containing
fully formed merozoites) without merosome formation
regardless of maturity. Sometimes it was found as early as
42 h after sporozoite infection, hours before merozoite
differentiation begins. The end result of this alternative
process was the formation of large faintly fluorescent EEF
ghosts containin g some cellular debris and a few dead
merozoites. We interpret this rapid conversion of EEFs to
ghosts as abortive liver stage development.
Merozoite content of EEFs has historically been difficult to
estimate due to the large number of parasites and their high
packing density. Based on measurements of the size and
merozoite content of small merosomes combined with size
measurements of EEFs and an appropriate mathematical
algorithm [42], we were able to calculate the number of
merozoites in an EEF (Table S2). Under intravital imaging
conditions, mature P. yoelii EEFs measured 40–75 lm and the
calculated space effectively occupied by a merozoite is a
sphere of 2.2 lm diameter. Using this effective size, we
calculated that individual P. yoelii sporozoites produce
roughly 4,200–29,000 merozoites per EEF. This number is in
general agreement with older estimates of EEF merozoite
content [5,12,22,23,52–57] (Table S1). An exception is P.
falciparum, which produces considerably larger numbers of
hepatic merozoites, most likely because of the small size of
the parasites. As far as we know, our analysis of the number of
merozoites produced in hepatocytes is the first such analysis
based on actual merozoite counts and host cell measure-
ments. Precision is limited by variations in measurements, but
basing calculations on direct in vivo measurements enhances
accuracy.
Earlier studies conducted by us and others had suggested
that merosome budding may precede completion of mer-
ozoite differentiation [14,15]. One factor that helped lead to
this interpretation is that GFP expressed in the parasite
stroma can obscure the parasites in mature EEFs. We now
show that prior to merosome formation, the signal of the
stromal GFP fluorescence equaled that of the merozoite
cytoplasm, thus preventing clear definition of parasites
enmeshed in the stroma. At the onset of merosome budding,
the stromal GFP emission signal decreased abruptly thus
revealing the presence of the already formed fluorescent
parasites (Figure 3A–3E and Video 5). Two factors contribute
to this reduction in fluorescence of material surrounding the
parasites: dilution and loss of cytosolic GFP. Dilution of GFP
results from PV disassembly and mixing of fluorescent
parasite stroma with non-fluorescent host cytoplasm. Loss
of GFP is caused by leakage of the fluorochrome into the
environment. In agreement with reports that the hepatocyte
membrane becomes permeable at late stages of infection with
P. berghei [5], we found that merosome-forming EEFs are
typically surrounded by a halo of gr een fluorescence.
Optimization of the imaging conditions allowed u s to
visualize the parasites inside mature EEFs and revealed that
merosomes always contain mature merozoites. Thus, mer-
ozoites maturation precedes merosome formation.
Depending on the approach used for measurement, the
reported diameters of hepatic and pulmonary capillaries vary
greatly. For example, when measured in perfusion-fixed liver
tissue, the sinusoidal diameter ranged from 4–6 lm to 9–12
lm [58–60]. A crucially important factor is the pressure
applied during perfusion fixation, because the sinusoidal
diameter is known to vary with changes in blood pressure
[61,62]. To determine the sinusoidal diameter under normal
blood pressure conditions, we used live Tie2-GFP mice [32],
whose fluorescent endothelia clearly delineate the boundaries
of the sinusoidal lumen [31]. In agreement with earlier in vivo
microscopic studies, which reported a diameter of 6 lm for
portal sinusoids and 7 lm for central sinusoids [58], we found
by intravital imaging that liver sinusoids measure 6.7 6 1.9
lm in diameter. Similar differences between fixed and live
specimens were reported for the size of alveolar capillaries.
While vascular casts of the lung suggested that alveolar
capillaries measure 6.69 6 1.39 lm in diameter [63], intravital
measurements determined a functional diameter of only 1–4
lm [64,65]. Regardless which liver sinusoid and lung capillary
measurements are relied upon and regardless of the drastic
reduction in merosome size after leaving the liver, mero-
somes still exceed the size of the lumen of the micro-
vasculature of both liver and lung. Since even the largest
merosomes were eventually transported out of the liver, the
much smaller extrahepatic merosomes would be expected to
be malleable enough to be able to pass though the pulmonary
capillary bed. Therefore it is somewhat surprising that the
lungs effectively clear the blood of all merosomes, so virtually
none were detectable in arterial blood harvested from the left
ventricle, in the capillary beds of spleen, brain and kidney, or
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Plasmodium Merozoite Release in the Lung

in tail vein blood. The fact that the velocity in pulmonary
capillaries is somewhat higher than hepatic sinusoids [66–69]
makes this more unexpected. Consequently, the possibility of
a receptor-mediated mechanism for pulmonary merosome
arrest cannot be excluded.
Materials and Methods
Parasites. Anopheles stephensi mosquitoes were used to propagate
wild-type P. yoelii (strain 17 XNL) or PyGFP [14,70]. Sporozoites were
purified from the salivary glands of female A. stephensi mosquitoes
[71].
Animals. Mice were (1) Balb/c (Taconic Farms, Incorporated), (2)
Swiss Webster (Taconic Farms, Incorporated), or (3) Tie2-GFP mice, a
transgenic strain that expresses GFP in vascular endothelial cells
under control of the Tie2 promoter (STOCK Tg(TIE2GFP)287Sato/J;
Jackson Laboratory) [31,32]. Animals were maintained and used in
accordance with recommendations in the guide for the Care and Use
of Laboratory Animals.
Surgery for intravital microscopy. Mice were inoculated into the
tail vein with 0.3–1.5 3 10
6
PyGFP sporozoites. At 30–66 h p.i., the
animals were surgically prepared for intravital imaging of liver and
spleen as described [31] and anesthetized by intraperitoneal injection
of a cocktail of 50 mg/kg ketamine (Ketaset, Fort Dodge Animal
Health), 10 mg/kg xylazine (Rompun, Bayer), and 1.7 mg/kg
acepromazine (Boehringer Ingelheim Vetmedica). Reinjection of
the anesthetics at 30-min intervals allowed intravital microscopic
examination of the animals for at least 3 h [31].
Intravital confocal microscopy. After surgical preparation for
intravital imaging, mice were placed onto the stage of an inverted
Zeiss DMIRE2 microscope, equipped with a temperature-controlled
Ludin chamber, and analyzed with a Leica TCS SP2 AOBS confocal
microscope. Appropriate laser lines were used to excite GFP, various
other fluorochromes, and the natural autofluorescence of the mouse
tissues. Laser power was reduced to a minimum to avoid photo-
toxicity and bleaching. These optimized conditions allowed contin-
uous scanning of live PyGFP for a period of up to 6 h without any
apparent effect on viability. To assess parasite and host cell viability,
some mice were i.v. injected with 1–2 lg/ml of the membrane-
permeable nuclear dye Hoechst 33342 prior to confocal microscopy.
Other mice received 1 lg/ml PI in addition to detect dead host cells
and/or parasites.
Infection with P. yoelii. Mice were intravenously inoculated with 3
310
6
purified wt P. yoelii or 1 3 10
6
PyGFP salivary gland sporozoites
and various organs were removed at 52 h after infection. Tissue slices
were snap-frozen in liquid nitrogen or fixed with PBS containing 4%
paraformaldehyde for immunofluorescence labeling of cryosections
and with PBS containing 4% paraformaldehyde and 1% glutaralde-
hyde for electron microscopic examination [72,73].
Ex vivo organ analysis. At 30–66 h after infection with PyGFP,
major organs such as spleen, brain, kidney, or lung were removed,
placed into glass-bottom dishes, and kept moist with medium for
confocal microscopy analysis.
Thick blood smears. Blood was harvested from (1) the terminal
hepatic vein, (2) the aorta, or (3) a tail vein. To increase the
probability of detection, ten aliquots of 5 ll blood from each of these
sites were spread over an area of 1 cm
2
, allowed to dry, and stained
with Giemsa without prior fixation. Merosomes were counted and
expressed as average number 6 STD. In parallel, the number of
merozoites per merosome was determined accordingly.
Analysis of merosome infectivity. Two days after infection with 1.5
3 10
6
wt P. yoelii sporozoites, hepatic effluent and tail vein blood was
harvested from the same animal and parasitemia and merosome
content were deter mined using thin and thick blood smears,
respectively. 20-ll hepatic effluent, containing 1 3 10
5
infected
erythrocytes plus 167 merosomes, or tail vein blood containing the
same number of infected erythrocytes but no merosomes, was
intravenously inoculated into Swiss Webster mice (three mice per
group) and the parasitemia was monitored daily by Giemsa staining.
Liver perfusion and merosome immobilization. To improve the
recovery of parasite material from the l iver, merosome s were
dislodged from hepatic sinusoids by perfusing mouse livers via the
portal vein with oxygenated medium at 5 ml/min for 10–30 min. The
effluent was collected in two fractions: fraction 1 was collected from
the Vena cava inferior and contained mainly red blood cells; fraction
2 was collected from the Vena cava superior after ligation of the Vena
cava inferior. The cells were washed and allowed to settle onto cover
slips or glass-bottom dishes (WillCo Wells) treated with Alcian blue
[74] for live cell imaging. Nuclei of merosomes were visualized with
the membrane permeable nucleic acid stains Hoechst 33342 (1–2 lg/
ml) or SYTO-64. Nuclei of dead parasites were determined with
membrane impermeable PI (1 lg/ml). Merosome membranes were
stained with 5 lg/ml FM 4–64 FX (Molecular Probes). Annexin V
Alexa Fluor 488 conjugate or YO-PRO-1 (0.1 lM) were used to detect
evidence of programmed cell death in live merosomes. Tissue
sections were stained with a BrdU TUNEL assay kit (Molecular
Probes) according to manufacturer’s guidelines.
Merozoite counting. Alcian blue–immobilized PyGFP merosomes
were fixed and labeled with the red nuclear dye SYTO-64. 3-D stacks
were scanned by confocal microscopy and the number of merozoite
nuclei was counted using a 3-D object count plug-in of ImageJ (NIH
freeware). Merozoite number and merosome diameter were then
entered into a formula for efficient packing of equal small spheres in
a large sphere ( n ¼ 0.7405 [1–2D] / D
3
þ 1 / [2D
2
]; D ¼ d
merozoite
/d
liver
stage
) [42] to determine the effective diameter/volume merozoites
occupy inside merosomes. Based on these calculations and the
diameter of PyGFP liver stages measured by intravital microscopy,
the merozoite content of P. yoelii liver stages was estimated in relation
to size.
Immunofluorescence microscopy. Frozen sections of 10-lm thick-
ness were prepared with a Reichert-Jung Frigocut cryostat. Parasites
were labeled with a mAb directed against the P. yoelii merozoite
surface protein MSP-1, a kind gift from W. Bergman [75]. A rabbit
antiserum, which was originally generated against the PVM-associ-
ated protein from P. berghei, but exhibits cross-reactivity with P. yoelii
UIS4 [38,76], was used to label the PV in P. yoelii–infected hepatocytes.
Affinity-purified goat IgG against the murine asialoglycoprotein
receptor ASGR1 was from R&D Systems. Incubation with the primary
antibodies was followed with protein A conjugated to fluorescein
isothiocyanate (PA-FITC; Molecular Probes), anti-goat IgG conju-
gated to Texas Red (GAR-TR; Molecular Probes), or goat anti-rabbit
IgG conjugated to Texas Red (GAM-TX; Molecular Probes) in color-
matching fluorochrome combinations. In case of a single FITC label,
the specimens were counterstained with 0.1% Evans blue in PBS.
Immunofluorescence-labeled frozen tissue sections were examined by
confocal microscopy.
Transmission electron microscopy. Mouse liver or lung tissue was
fixed with 1% glutaraldehyde and 4% paraformaldehyde in PBS,
post-fixed with 1% osmium tetroxide and 1.5% potassium hexacya-
noferrate, stained en bloc with 1% uranyl acetate, dehydrated in
ethanol, and embedded in Epon as described [72,73]. Semithin
sections were cut with an RMC MT-7 ultramicrotome and photo-
graphs were taken with Kodak Ektachrome 160T slide film using a
Nikon FX-35DX/UFX-DX camera/exposure system. Thin sections
were post-stained with uranyl acetate and lead citrate and viewed
with a Zeiss EM 910 electron microscope [73].
Image processing. Electron microscopy negatives and Ektachrome
slides were scanned with a Hewlett Packard Scanjet 5370C. All digital,
electron, or confocal microscopy images were processed using Image-
Pro Plus (Media Cybernetics), Adobe Photoshop (Adobe), and
AutoDeBlur (AutoQuant Imaging, Incorporated) software.
Supporting Information
Figure S1. Infiltration of Disintegrating EEFs by Inflammatory Cells
Leads to Granuloma Formation
(A) Semithin Epon section showing a granuloma (outlined by arrows)
in the liver of a mouse 52 h after infection with P. yoelii. Inflammatory
cells have infiltrated the remnants of an EEF. The surrounding tissue
shows a normal arrangement of hepatocytes and sinusoids. Bar ¼ 10
lm.
(B) Intravital confocal scan of mouse liver showing an EEF in the
process of disintegration. Only a small amount of merozoites (green)
is left in the remains of the EEF. Inflammatory cells that have
infiltrated the remains of the former EEF are visualized by nuclear
staining with Hoechst (blue). Bar ¼ 10 lm.
(C) Electron microscopy identifies the inflammatory cells as mono-
nuclear phagocytes and polymorph nuclear granulocytes. Parasite
remains can be detected inside phagolysosomes (arrows) of these
phagocytes. Mo, mononuclear phagocytes; PMN, polymorph nuclear
granulocytes. Bar ¼ 5 lm.
(D) Part of a granuloma showing necrotic remnant bodies and
merozoites () embedded in the loose matrix of a degenerating EEF
(arrows). It appears that some parasite material has discharged
(arrowheads) i nto the space between t he fo rmer EEF and a
neighboring hepatocyte. RB, remnant body; H, hepatocyte. Bar ¼ 5
lm.
PLoS Pathogens | www.plospathogens.org November 2007 | Volume 3 | Issue 11 | e1711665
Plasmodium Merozoite Release in the Lung

Found at doi:10.1371/journal.ppat.0030171.sg001 (10.9 MB TIF).
Figure S2. Ghost Formation after EEF Decay
Individual frames from an intravital video demonstrating the events
involved in rapid EEF decay.
(A and B) At the onset of disintegration, green fluorescent parasite
content streams out of an immature EEF into an adjacent sinusoid
(arrow).
(C) Eventually, a large faintly fluorescent ghost is left behind. Note
that the membrane of the EEF never lost contact with neighboring
cells.
(D) Decay of a mature EEF. The sudden loss of GFP from the parasite
cytosol reveals the presence of large numbers of merozoites (arrow-
heads; see Video S9). Bars ¼ 10 lm.
Found at doi:10.1371/journal.ppat.0030171.sg002 (6.2 MB TIF).
Figure S3. Liver Perfusion Enhances Merosome Formation
(A) Ex vivo confocal analysis of a non-perfused mouse liver. The
presence of oversized PyGFP merosomes suggests that the absence of
blood flow prevented them from subdividing into smaller units. Bar ¼
20 lm.
(B) Ex vivo confocal analysis of a perfused mouse liver. Merosomes
extend over hundreds of micrometers from the EEF into various
sinusoids indicating that liver perfusion augmented me rosome
budding and separation from the host cell. Bar ¼ 100 lm.
Found at doi:10.1371/journal.ppat.0030171.sg003 (7.2 MB TIF).
Table S1. Size and Merozoite Content of Plasmodium EEFs
Comparison of the estimated numbers of merozoites produced by
various Plasmodium species in different hosts with the calculated data
obtained from P. yoelii–infected mice presented in this paper.
Found at doi:10.1371/journal.ppat.0030171.st001 (62 KB DOC).
Table S2. Merozoite Content of P. yoelii Merosomes and EEFs
Merosomes were collected from the hepatic effluent. A subpopula-
tion of small merosomes was used to measure the merosome diameter
in relation to the merozoite content. An algorithm for optimal sphere
packing (see Materials and Methods) was then used to calculate the
effective merozoite diameter in these merosomes. This effective
diameter (2.175 lm) was then entered into the same algorithm to
determine the number of merozoites contained in the much larger
EEFs in relation to their diameter.
Found at doi:10.1371/journal.ppat.0030171.st002 (35 KB DOC).
Video S1. Merozoite Release by Merosome Formation
Intravital confocal microscopy showing a medium-sized merosome in
the process of budding from an EEF (located outside of the optical
plane and therefore not shown) at 52 h after intravenous infection
with PyGFP sporozoites. The merosome cont ains hundreds of
merozoites and several non-fluorescent remnant bodies. The video
was recorded over a total period of 25 min. During this time, the
merosome subdivides into two smaller units.
Found at doi:10.1371/journal.ppat.0030171.sv001 (9.8 MB AVI).
Video S2. Merosome Budding
Shows budding of a large merosome from a mature EEF. Note the
movement within the EEF during the budding process. A few
individual merozoites are located in the vicinity of the infected
hepatocyte. Shows intravital confocal microscopy of a mouse liver 48
h after intravenous infection with PyGFP sporozoites. Total record-
ing time: 2 min and 20 s.
Found at doi:10.1371/journal.ppat.0030171.sv002 (9.8 MB AVI).
Video S3. Merozoite Release by Merosome Budding
An EEF releases merozoite-filled merosomes into the bloodstream.
Note the ragged surface of the EEF, which is gradually decreasing in
size during the budding process. Total recording time: 2 min and 36 s.
Found at doi:10.1371/journal.ppat.0030171.sv003 (9.4 MB AVI).
Video S4. Individual Merozoites Are Released from a Disintegrating
EEF
At 54 h after intravenous infection with PyGFP sporozoites, a
disintegrating mature EEF can be seen slowly discharging individual
merozoites into the bloodstream. Total recording time: 2 min and
36 s.
Found at doi:10.1371/journal.ppat.0030171.sv004 (9.7 MB AVI).
Video S5. Rapid EEF Decay Leading to Ghost Formation
The video shows the initial stage of EEF ghost formation. The mature
EEF suddenly leaks GFP into the environment. At the same time, the
GFP emission signal in the EEF stroma decreases revealing the
presence of parasites. Shows mouse liver 45 h after intravenous
infection with PyGFP sporozoites. Total recording time: 6 min and 31 s.
Found at doi:10.1371/journal.ppat.0030171.sv005 (9.5 MB AVI).
Video S6. Fast Merosome Transport along a Liver Sinusoid
Multiple merosomes are transported towards the central vein. The
merosomes bud from an EEF, which is located outside of the focal
plane and therefore not visible, and contain mature merozoites and
non-fluorescent remnant bodies. Several other merosomes remain
stationary during the observation period. The video shows intravital
confocal microscopy of a mouse liver 52 h after intravenous infection
with PyGFP sporozoites. Total recording time: 7 min and 11 s.
Found at doi:10.1371/journal.ppat.0030171.sv006 (9.7 MB AVI).
Video S7. Slow Transport of Large Merosomes toward the Central
Vein
Two large merosomes (on the left) are transported along a liver
sinusoid, are temporarily arrested at a sinusoidal bifurcation, and
reverse direction of movement. The resulting obstruction of the
blood flow prevents another merosome (on the right) from entering
the same sinusoid. Note the high malleability of the merosomes. The
video shows intravital confocal microscopy of a mouse liver 52 h after
intravenous sporozoites infection with PyGFP. Total recording time:
5 min and 23 s.
Found at doi:10.1371/journal.ppat.0030171.sv007 (9.5 MB AVI).
Video S8. Subdivision of a Larger Merosome into Smaller Units
While arrested at a sinusoidal bifurcation, a medium-sized merosome
releases a smaller unit which is rapidly swept away by the blood-
stream. Note the large number of mature merozoites inside the
merosome. The video shows intravital confocal microscopy of a
mouse liver 52 h after intravenous infection with PyGFP sporozoites.
Total recording time: 2 min and 36 s.
Found at doi:10.1371/journal.ppat.0030171.sv008 (9.4 MB AVI).
Video S9. Newly Released Merosomes Are Large Branched Structures
The animated 3-D stack generated from a perfused mouse liver 52 h
after infection demonstrates all aspects of merosome release. Ex vivo
confocal analysis shows an EEF (center) that has released numerous
merosomes of various sizes into adjacent sinusoids. Note that the
largest merosomes are branched and contain hundreds of merozoites.
Found at doi:10.1371/journal.ppat.0030171.sv009 (10.1 MB AVI).
Video S10. Merozoite Release into the Lung Microvasculature
Ex vivo confocal microscopy shows a group of individual merozoites
being released from a pulmonary merosome (on the left). Note the
mobility of the free merozoites. Shows mouse lung 52 h after
intravenous infection with PyGFP sporozoites. Total recording time:
1 min and 18 s.
Found at doi:10.1371/journal.ppat.0030171.sv010 (9.4 MB AVI).
Acknowledgments
Many thanks to Drs. Jerome Vanderberg and Allen B. Clarkson for
critically reviewing the manuscript. We are indebted to Dr. William
Bergman (Dre xel Un iversity College of Medicine, Philadelphi a,
Pennsylvania) for a kind gift of a monoclonal antibody against P.
yoelii MSP- 1 and to Dr. T om Cameron, Skirbal l Institute for
Biomolecular Medicine, New York University, for helpful suggestions
regarding the in vivo use of PI.
Author contributions. KB, CK, and UF conceived and designed the
experiments, performed the experiments, analyzed the data, and
wrote the paper. SHIK contributed reagents/materials/analysis tools.
TS sponsored KB’s dissertation.
Funding. This work was supported by National Institutes of Health
grants RO1 AI51656 and S10 RR019288 and National Science
Foundation grant 9977430 to UF and a grant from the Ellison
Medical Foundation and a Seattle Biomedical Research Institute
Innovation Grant to SHIK. KB received a pre-doctoral fellowship
from the Karl-Enigk-Stiftung, Hannover, Germany.
Competing interests. The authors have declared that no competing
interests exist.
PLoS Pathogens | www.plospathogens.org November 2007 | Volume 3 | Issue 11 | e1711666
Plasmodium Merozoite Release in the Lung

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PLoS Pathogens | www.plospathogens.org November 2007 | Volume 3 | Issue 11 | e1711668
Plasmodium Merozoite Release in the Lung