Beyond the microcirculation: sequestration of infected red blood cells and reduced flow in large draining veins in experimental cerebral malaria

Oelschlegel, A. M.; Bhattacharjee, R.; Wenk, P.; Harit, K.; Rothkötter, H-J; Koch, S. P.; Boehm-Sturm, P.; Matuschewski, K.; Budinger, E.; Schlüter, D.; Goldschmidt, J.; Nishanth, G.

doi:10.1038/s41467-024-46617-w

Download PDF

Article
Open access
Published: 16 March 2024

Beyond the microcirculation: sequestration of infected red blood cells and reduced flow in large draining veins in experimental cerebral malaria

Nature Communications volume 15, Article number: 2396 (2024) Cite this article

1248 Accesses
16 Altmetric
Metrics details

Subjects

Abstract

Sequestration of infected red blood cells (iRBCs) in the microcirculation is a hallmark of cerebral malaria (CM) in post-mortem human brains. It remains controversial how this might be linked to the different disease manifestations, in particular brain swelling leading to brain herniation and death. The main hypotheses focus on iRBC-triggered inflammation and mechanical obstruction of blood flow. Here, we test these hypotheses using murine models of experimental CM (ECM), SPECT-imaging of radiolabeled iRBCs and cerebral perfusion, MR-angiography, q-PCR, and immunohistochemistry. We show that iRBC accumulation and reduced flow precede inflammation. Unexpectedly, we find that iRBCs accumulate not only in the microcirculation but also in large draining veins and sinuses, particularly at the rostral confluence. We identify two parallel venous streams from the superior sagittal sinus that open into the rostral rhinal veins and are partially connected to infected skull bone marrow. The flow in these vessels is reduced early, and the spatial patterns of pathology correspond to venous drainage territories. Our data suggest that venous efflux reductions downstream of the microcirculation are causally linked to ECM pathology, and that the different spatiotemporal patterns of edema development in mice and humans could be related to anatomical differences in venous anatomy.

Brain endothelial GSDMD activation mediates inflammatory BBB breakdown

Article 17 April 2024

Disease-associated astrocyte epigenetic memory promotes CNS pathology

Article 20 March 2024

Blood–brain barrier disruption and sustained systemic inflammation in individuals with long COVID-associated cognitive impairment

Article Open access 22 February 2024

Introduction

Cerebral malaria (CM) is a severe neurological complication of human malaria mostly caused by the parasite Plasmodium falciparum (Pf). It is the most common non-traumatic encephalopathy in the world with a high mortality rate. Globally, an estimated number of 608,000 patients died of malaria in 2022, while 96% of the malaria-related deaths occurred in sub-Saharan Africa, primarily in infants and children below the age of 5 years (WHO, 2023¹).

Patients with acute infection can present with a diffuse CM encephalopathy, a rapid progressive coma, and seizures without return to consciousness^2,3. Magnetic resonance imaging (MRI) findings in children in sub-Saharan Africa indicate that increased brain volume and brain swelling are hallmarks of the disease, and patients die from brain herniation^4,5. MRI measures of brain edema strongly correlate with the outcome in pediatric CM⁶.

The key pathological findings in post-mortem human brains are intravascular sequestration of and congestion with infected red blood cells (iRBCs)^7,8,9,10,11. In contrast to other parasitic diseases of the CNS, Pf-infected red blood cells in CM remain primarily intravascular^2,3,12,13. However, in murine experimental CM (ECM) some iRBCs have also been observed perivascularly in the parenchyma¹⁴. The cascade of events linking infection and brain swelling has remained elusive and is much debated^4,15. Theories focus either on a cytokine storm triggering inflammation and affecting vascular integrity or mechanical obstructions of cerebral blood flow (CBF) due to iRBC sequestration in the microcirculation^2,3,12,13. The former is primarily based on ECM models, while the latter originates from post-mortem histology of CM patients. While the relevance of ECM to CM continues to be viewed differently^16,17, there is growing consensus that studies with experimental models of CM are central to better define pathogenesis and identify early signatures that distinguish disease progression from parasite propagation^14,18. One central controversy is to which extent iRBC sequestration in cerebral microcapillaries, a distinctive hallmark of CM, also contributes to the disease in ECM^{12,19,20,21,22,23,24}. Therefore, insights into the temporal development of disease events in combination with their regional distribution might provide a better understanding of the pathophysiological processes leading to CM.

As the spatial patterns of brain swelling differ in different subgroups of patients⁴, it could be highly informative to correlate at, or close to, the onset of cerebral edema, the spatial patterns of iRBC sequestration with spatial patterns of edema development and potential perfusion deficits, which are to be expected upon mechanical obstruction. However, such studies are hardly feasible in humans, especially in the most affected countries with limited access to the imaging technologies needed.

Herein, we address these questions in murine ECM focusing on Plasmodium berghei ANKA (PbA) infected C57BL/6 wild-type (wt) mice. We extend parts of our study to murine malaria models that do not develop ECM or brain pathology. After studying brain iRBC accumulation in infected mice using quantitative real-time-PCR (q-PCR), we use single-photon emission computed tomography (SPECT) for imaging in vivo the distribution of ^99mTechnetium (^99mTc)-labeled iRBCs in the brain at day 5 post-infection (p.i.), an early stage of the disease when first clinical symptoms start appearing. We combine this approach with T2-weighted (T2w) MRI for edema monitoring as well as SPECT-imaging of CBF and MR-angiography. These imaging approaches allow us to detect potential sequelae of iRBC sequestration on cerebral perfusion at different spatial levels, i.e., capillary flow and large draining veins. We also relate these data to histological data at different time points after infection and to the expression of inflammation markers, as studied by q-PCR and immunohistochemistry, with and without anti-malarial treatment.

Results

Early-stage accumulation of PbA-infected red blood cells in large cerebral draining veins in C57BL/6 wt mice

We initiated our study by q-PCR analysis and quantified the PbA-specific gene cytochrome B in the brains of infected C57BL/6 wt mice at days 5 and 7 p.i. and, as control, in uninfected mice. Abundance of parasite mRNA was quantified in three different cerebral regions, olfactory bulb, cerebral cortex, and brainstem (Fig. 1A). Interestingly, PbA cDNA could be detected as early as day 5 p.i., prior the onset of noticeable disease symptoms (Fig. S1). As the disease progressed until day 7 p.i., the parasite load increased in all three regions of the brain. Of note, the parasite burden remained significantly higher in the olfactory bulb compared to cerebral cortex and brainstem (Fig. 1A).

**Fig. 1: PbA-infected red blood cells accumulate at an early stage in large draining veins of C57BL/6 wild-type and C57BL/6 *prf*⁻^/⁻ mice, but not BALB/c mice.**

In order to study the spatial distribution of iRBCs at an early stage in more detail, we labeled PbA-infected RBCs with the gamma-emitter ^99mTc and studied their biodistribution in PbA-infected C57BL/6 wt mice in vivo using SPECT. 1 h after injection of the labeled iRBCs, we imaged the ^99mTc-distributions in the heads of uninfected and infected mice at day 5 p.i. An accompanying whole-body scan was used to precisely determine standardized uptake values (SUVs). SUVs represent the ratio of the ^99mTc-concentration in a voxel and the whole-body ^99mTc-concentration. Peak differences in ^99mTc-content between infected mice and controls were found in areas corresponding to large cerebral draining veins or sinuses, in particular the rostral rhinal veins, the superior sagittal sinus, the straight sinus, and the transverse sinuses (Fig. 1B–F). The distribution of the differences within the superior sagittal sinus was inhomogeneous, strongly favoring the T-shaped junctions at the rostral and caudal confluences of the sinuses (Fig. 1B–F). An additional lower-intensity peak was found in the middle of the sinus (Fig. 1B–D). Literature data suggest that this peak could correspond to an entry site of larger draining veins²⁵.

We then imaged, as further controls, the iRBC distribution in PbA-infected perforin-deficient mice (C57BL/6 prf⁻^/⁻) (Fig. 1G–I) and BALB/c mice (Fig. 1J, K). In both C57BL/6 prf ⁻^/⁻ and BALB/c mice PbA replicates in erythrocytes and causes anemia, but not ECM^26,27. The spatial distribution of the iRBCs in C57BL/6 prf⁻^/⁻ mice closely mimicked that of the C57BL/6 wt mice (Fig. 1G–I), but the ratio of iRBCs in infected compared to non-infected mice was lower in C57BL/6 prf⁻^/⁻ mice (Fig. 1L). No differences were found between infected and uninfected BALB/c mice (Fig. 1J–L). The course of parasitemia was comparable between the infected C57Bl/6 wt mice and BALB/c mice (Fig. S2B), indicating that the genetic background of the host determines the outcome of the vascular pathogenesis in ECM.

Together, our data on the distribution of ^99mTc-labeled iRBCs identified accumulation in large cerebral draining veins at an early time point prior to clinical signs of progression to ECM. In infection models that progress to malaria-related anemia instead of ECM this accumulation was either less pronounced (C57BL/6 prf⁻^/⁻ mice) or absent (BALB/c mice).

Reduced flow in large draining cerebral veins in PbA-infected mice

Our iRBC-SPECT findings suggested that venous efflux in large draining veins might be impaired already at an early stage of disease onset indicative of a trajectory towards ECM. We tested this hypothesis using time of flight MR-angiography (TOF-MRA) with sequences for imaging blood flow in caudo-rostral direction in C57BL/6 wt, C57BL/6 prf⁻^/⁻ and BALB/c mice (Fig. 2). We used a novel automated workflow atlas registration described previously by Koch and colleagues²⁸ for aligning all MR-angiographies in the same anatomical reference frame. We adopted this workflow to the present ECM study, which greatly facilitated group-level analyses at single-vessel levels (Fig. 2).

**Fig. 2: Magnetic resonance angiography uncovers reduced flow in large draining veins only in PbA-infected mice on trajectory to ECM.**

For quantitative analysis, we measured the signal intensities in a volume-of-interest (VOI) segmented from the superior sagittal sinus (sss) and the rostral rhinal vein (rrv) of uninfected control mice. In this VOI, the mean signal intensity dropped by 22% in infected C57BL/6 wt mice at day 5 p.i. compared to controls (p < 0.01), and further dropped by 39% at day 7 p.i., as compared to controls (p < 0.001) (Fig. 2). Peak intensities inside the sss and rrv dropped by more than 70% at day 7 p.i. compared to the uninfected controls. The variance at day 7 p.i. was the highest of all groups measured, and in some individuals the flow was barely detectable.

In our anatomical T2w images (Figs. 2 and 3) we found prominent signal increases indicating severe edema at day 7 p.i. in the same structures as described previously²⁹, in particular in white matter fiber tracts, such as the corpus callosum (Figs. 2 and 3). At day 5 p.i. we were unable to unambiguously identify or exclude signs of edema. In line with a previous study³⁰, we found no evidence for edema in infected C57BL/6 prf⁻^/⁻ mice. Although mean signal intensity in the sss-rrv VOI decreased slightly, the decreases did not pass thresholds of significance (Fig. 2). In infected BALB/c mice, we detected a slight, albeit non-significant, increase in mean signal intensity (Fig. 2). This might simply reflect noise but could also be related to blood flow alterations as a consequence of the systemic infection.

**Fig. 3: In PbA-infected C57BL/6 wt mice flow is reduced most severely in the superior sagittal sinus and the rostral rhinal vein.**

In all groups of mice, we noted that the sss gives of a branch that runs above this sinus into the rrv (Fig. 3 d0 Bregma +3.0 and +0.8; Fig. 2, BALB/c mice d0). The mean signal intensity in all groups was higher in this upper branch as compared to the lower (Fig. 2).

We conclude from the TOF-MRA data that in PbA-infected C57BL/6 wt mice a reduction in venous efflux precedes edema formation and progresses rapidly, corresponding to the severity of the disease, as quantified by clinical scores (Fig. S1).

Histology confirms the presence of iRBCs in large draining veins of PbA-infected C57BL/6 wt mice

We then looked for vascular pathological alterations in the large draining veins and studied PbA-infected C57BL/6 mice histologically (Fig. 4). We reasoned that in order to not miss pathological intravascular phenomena, for instance rosetting, we should omit transcardial perfusion. Our analysis was based on immersion-fixed decalcified material.

**Fig. 4: Large deposits accumulate in iRBCs in draining veins at day 7 after PbA-infection in C57BL/6 wt but not in C57BL/6 *prf*⁻^/⁻ mice.**

At day 7 p.i., when the mice showed typical symptoms of ECM with a mean RMCBS score of 8, we found iRBCs with dark black-brown deposits with diameters of several micrometers most likely representing hemozoin in the large draining veins of PbA-infected C57BL/6 wt mice (Fig. 4A–J). IRBCs were also present in the large draining veins of PbA-infected C57BL/6 prf⁻^/⁻ mice, but we were unable to find the large-diameter deposits that characterized the C57BL/6 wt iRBCs (Fig. 4K–U). To further clarify whether the presence of these deposits might be related to or correlate with the development of the ECM pathology we included in our analysis PbNK65-infected C57BL/6 wt mice that are known to not develop cerebral malaria²⁷. Here, as studied on day 7 p.i., we also did not find the black-brown deposits present in the iRBCs of PbA-infected C57BL/6 wt mice (Fig. 2S2A–E). In all three models we found black-brown deposits in bone marrow cells arguing for skull bone marrow infections (Fig. 4J, T, S2C). Bone marrow infections in murine models have been described earlier³¹. The infection of the murine skull bone marrow, however, is to the best of our knowledge a novel finding.

At day 5 p.i., which was the earliest time point examined, we found in PbA-infected C57BL/6 wt mice iRBCs in the large draining veins, either single iRBCs among non-infected RBCs (Fig. 5M, N, O) or densely packed groups of iRBCs (Fig. 5J, K, L) and also iRBCs in direct contact to the vessel walls (Fig. 5E, F, G). The same types of deposits that we detected in these iRBCs, i.e., dark-brown granules that polarized light, were also found in parts of the vessel walls (Fig. 5F, G) suggesting vessel reactions and sequestration of iRBCs in the large draining veins, or certain compartments of these veins (Fig. 5C).

**Fig. 5: At day 5 p.i. large draining veins in C57BL/6 wt mice show vessel wall reactions and sequestration of iRBCs.**

In conclusion, our histological results confirm the findings of our iRBC-SPECT-imaging study. IRBCs are present in large draining veins, and vessel wall reactions are indicative of iRBC sequestration.

MR-angiography and histology uncover two parallel venous streams from the superior sagittal sinus to the rostral rhinal veins

In accordance with our TOF-MRA findings we also noted in our histological study that the sss, at around Bregma +0.6 mm splits into two vessels positioned on top of each other. Histologically and in X-ray CT images we find that the upper vessel runs within the diploic space and rostrally divides into left and right rrv (Fig. 6A–H) which have direct connections to the bone marrow (Fig. 6J). The lower vessel can be seen as the superior sagittal sinus proper as it runs below the bone into the rostral confluence of the sinus. Here, both venous streams are connected again (Fig. 6G). The sagittal TOF-MRA images (Fig. 2, BALB/c d0) in conjunction with a low-density area in the CT (Fig. 6E) indicate that a third connection exists between both streams in the caudal one-third of both vessels. In particular in the upper stream (Fig. 5C; 6P, Q, S, T) but also in the lower (Fig. S2F) we find vessel wall reactions in infected C57BL/6 wt mice.

**Fig. 6: Topographical relationships of major sites of pathology, the superior sagittal sinus and the rostral rhinal vein.**

Our combined histological, X-ray CT, and TOF-MRA study thus uncovered a vascular anatomy of the rostral superior sagittal sinus that is more complex than the traditional scheme. Two interconnected parallel venous streams instead of one single sinus run rostrally towards the rostral confluence of the sinuses and the rostral rhinal veins.

Impaired perfusion in territories of large draining cerebral veins in PbA- infected mice

In order to study the impact of early iRBC sequestration in the brain and the large draining veins on cerebral perfusion, we used ^99mTc-hexamethylpropylene amine oxime (^99mTc-HMPAO) SPECT, a technique widely used in clinical routine for imaging spatial patterns of CBF. We imaged uninfected and infected C57BL/6 wt mice at day 5 p.i. Notably, we did not find in any individual perfusion deficits indicating arterial infarctions, i.e., severely reduced flow in territories of the major arteries or branches thereof. Instead, we found a moderate decrease in cerebral perfusion, which was largely bilaterally symmetric and most prominent in the dorsocaudal olfactory bulb, the rostral prefrontal cortex, the caudal midline cortex, the adjacent visual cortices as well as in upper midline superior colliculus (Fig. 7). These regions are directly underlying or adjacent to the junctions of the sagittal sinus with the rostral rhinal veins and the transverse sinus, respectively, where the iRBC accumulation was most prominent (Fig. 1). This finding indicates early alterations in perfusion in territories of these large draining veins. We noted additional decreases in CBF in the hippocampal formation and entorhinal cortex (Fig. 7, Bregma −3.3), areas that finally drain into the straight sinus³². The tendency towards bilateral alterations would be in line with venous efflux reduction in an unpaired midline vessel playing a causal role. However, due to the small isolated foci of reduced CBF, the congruences of affected areas and venous drainage territories are less apparent than in the olfactory bulb/rostral rhinal vein region.

**Fig. 7: Cerebral blood flow is impaired early in territories of large draining veins.**

ECM-associated brain pathology and pro-inflammatory responses are late events in ECM pathogenesis

In order to relate the in vivo imaging findings to brain region-specific neuroinflammation and pathology, we studied the brains of C57BL/6 wt mice at corresponding time points using an array of histological and molecular techniques. Very few signs of brain pathology were visible at day 5 p.i. (Fig. 8A–F), which progressed until day 7 p.i., where the mice developed widespread neuroinflammation. Interestingly, the intracerebral infiltration of CD8⁺ T cells was negligible at day 5 p.i. and similar to uninfected mice (Fig. 8D). However, numbers of intracerebral CD8⁺ T cells were significantly increased at day 7 p.i. in all brain regions, predominantly in the sub-ventricular zone (36-fold) and midbrain region (33-fold) (Fig. 8D, Fig. S7). Thus, CD8⁺ T cells, which contribute unambiguously to the lethal outcome of ECM, increase only after the onset of iRBC accumulation and reduced venous efflux in the brain.

**Fig. 8: ECM-associated brain pathology progressively worsens and is not region-specific.**

This finding prompted us to analyze the spatial and temporal expression of intracerebral pro-inflammatory cytokines and chemokines in the olfactory bulb, cortex, and brainstem (Fig. 9). At day 5 p.i., when there was a significant accumulation of iRBCs in the brain (Fig. 1K), mRNA levels of the pro-inflammatory cytokines IFN-γ, and TNF and the chemokines CXCL9, CXCL10, CXCL11, which are essential for the recruitment of pathogenic CD8⁺ T cells to the brain, were only slightly induced. In marked contrast, at day 7 p.i. there was a significant increase in levels of the chemokines and cytokines throughout all three brain regions of PbA-infected mice (Fig. 9).

**Fig. 9: ECM augments cytokine and chemokine production in the later stages of the disease.**

In good agreement with the increase of CD8⁺ T cells and upregulation of cytokines and chemokines from day 5 to day 7 p.i., the numbers of GFAP⁺ astrocytes (Fig. 8A, S4) and Iba1⁺ microglia (Fig. 8B, S5) and CD31/PECAM-1⁺ staining intensity on endothelial cells (Fig. 8C, S6) increased in all brain regions gradually from day 0 to day 7 p.i. Of note, the strongest increase was again from day 5 to day 7 p.i. further illustrating that neuroinflammation and activation of brain resident cell populations developed later than significant accumulation of iRBCs in cerebral blood vessels. Interestingly, as observed in human CM, in our murine ECM model large hemorrhagic lesions in the brainstem were observed (Fig. 8F, S8), which also stained positive for cleaved caspase-3, an apoptosis-inducing effector caspase (Fig. 8E, S9), reproducing CM pathology in ECM. Hemorrhagic lesions and apoptotic cells emerged only later at day 7 but not at day 5 p.i.

Taken together, our data show that neuroinflammation is a late event and develops after significant accumulation of iRBCs in cerebral blood vessels in ECM.

Early anti-malaria treatment reverses brain pathology

Since our data support the notion that intracerebral iRBC accumulation is essential for ECM development, we next wanted to validate whether the pro-inflammatory response was a consequence of iRBC accumulation. To this end, we treated infected C57BL/6 wt mice with the anti-malarial drug pyrimethamine starting from day 5 p.i. Anti-malarial treatment rapidly reduced the amount of iRBC within 24 h (Fig. S10A) and resulted in survival of all mice (Fig. S10B). Furthermore, BBB damage in brains of pyrimethamine-treated mice was absent in contrast to untreated animals (Fig. S8C).

Importantly, treatment with the antimalarial drug reduced brain pathology (Fig. 8) and resulted in low pro-inflammatory cytokine and chemokine responses at day 7 p.i. (Fig. 9). These findings indicate that anti-malarial treatment at the time of intravascular accumulation of iRBC can prevent the complications leading to the pathogenesis of ECM suggesting that sequestration of infected red blood cells and reduced venous efflux precede inflammation in experimental cerebral malaria.

Disruption of the rostral migratory stream is a consequence of iRBC accumulation

We included in our study the analysis of the rostral migratory stream (RMS) and replicated the findings of Hoffmann et al. ²⁹. in C57BL/6 wt mice. The migration and proliferation of neuroblasts in the rostral migratory stream (RMS) were disturbed already at day 5 p.i. as indicated by reduction in the number of doublecortin (DCX) and bromodeoxyuridine (BrdU) stained neuroblasts (Fig. S3A, B). The disruption progresses further until day 7 p.i., where the RMS was completely disturbed.

Interestingly, early treatment not only prevented the damage of the RMS, but also fostered its repair, indicating that the disruption of the RMS is consequence of RBC accumulation (Fig. S3A–E).

Discussion

In the present study we analyzed, for the first time, the 3D in vivo distribution of iRBCs in experimental cerebral malaria at an early stage of the disease, where neurological symptoms, inflammation, and edema are virtually absent. Using SPECT-imaging in a murine ECM model we found significant increases of ^99mTc-labeled iRBCs in the brains and in large draining veins or sinuses. Simultaneously, venous efflux in large cerebral draining veins and sinuses, as determined by TOF-MRA, decreased at day 5 p.i. and was further reduced to nearly undetectable levels in symptomatic affected mice at day 7 p.i. SPECT-imaging of CBF using ^99mTc-HMPAO as a tracer revealed moderate decreases in perfusion at day 5 p.i. Affected regions were found in the caudal olfactory bulb, rostral prefrontal cortex as well as in the posterior cingulate and visual cortex, areas from which venous efflux drains into the rostral rhinal veins or the superior sagittal sinus, respectively^25,32,33.

The 3D distribution of iRBCs as determined with SPECT is in excellent agreement with previous 2D optical ex vivo whole-brain images of the parasite distribution, obtained with luciferase-expressing PbA^19,20,34. Although the relationship of the parasite distribution to the venous sinuses has remained unnoticed, the published data can now be readily interpreted as 2D projection images of the 3D distribution described in our study.

We independently confirmed the presence of iRBCs in the large draining veins histologically. We note that in a previous comprehensive study of the distribution of GFP-labeled parasites in the murine brain the sinuses were not included¹⁴. We propose that attention to parasite distribution to the venous sinuses and omission of transcardial perfusion will be important for future studies on the dynamics of ECM onset.

As early as day 5 p.i., we find in PbA-infected C57BL/6 wt mice not only inside the iRBCs but also in vessel walls black-brown deposits polarizing light and thus most likely representing hemozoin. This suggests a vascular or endothelial reaction to iRBCs as described in the literature³⁵ and iRBC sequestration. At day 7 p.i. we find large-diameter deposits similar in size and shape to those of rosetting iRBCs, as described before³⁶.

To substantiate our findings, we also included two murine systems, PbA infections in C57BL/6 prf⁻^/⁻ and BALB/c mice, which develop severe anemia after protracted parasitemia but do not develop ECM. Venous efflux was functional and large-diameter deposits were not detectable in these infection models. In agreement with earlier results from MR-imaging³⁰ infected C57BL/6 prf⁻^/⁻ mice did not develop edema. Our findings suggests that vessel pathology, as shown by deposition of parasitic material in the vessel wall, requires contribution of the immune system, likely by activating endothelial cells, a process in which perforin might be involved. We also note that edema was only detectable after flow in the large draining veins was substantially compromised.

Our interpretation is supported by a recent study³⁷ on protective effects of hypothyroidism in ECM. In hypothyroid C57BL/6 wt mice, PbA infections did not result in increased intracranial pressure and edema despite inflammation and BBB-breakdown. In these mice the flow in the large vessels, as determined with TOF-MRA, was maintained. Although angiographies were not analyzed at the single-vessel level, the flow in the sinuses is clearly visible in these TOF-MRA images³⁷.

Together, these findings challenge the traditional view of edema generation by pathological alterations in the microcirculation and are fully compatible with an alternative concept of edema generation as a consequence of downstream reductions of venous efflux. We propose that the spatiotemporal patterns of edema development, and of ECM pathology in general, are triggered by reduced efflux in the large draining veins and later exacerbated by local pathological alterations within the microcirculation.

In rodents, the superior sagittal sinus drains into the rostral rhinal veins^25,33 forming a major venous efflux route that differs from humans. In PbA-infected C57BL/6 mice, we find the highest intensity of labeled iRBCs at the T-shaped junction of the sagittal sinus and the left and right rostral rhinal veins. The earliest signs of pathological alterations occur in regions draining into these veins and earliest signs of blood-brain barrier disruptions are detectable in brain tissue directly below the junction. A previous MR-study showed the first signs of edema and blood-brain barrier disruption in the caudal olfactory bulb, a region also severely affected by hemorrhages at later stages²⁹. From there the edema spreads caudally along white matter fiber tracts affecting, among other structures, the corpus callosum. The spatiotemporal dynamics of edema onset and progression are unlikely to be influenced by host factors governing iRBC densities in the microcirculation. Instead, impaired flow in the superior sagittal sinus and the rostral rhinal veins provide a plausible explanation for edema dynamics and parasite distribution in the brain. According to this model, differences between mice and humans in the spatial patterns of edema onset and progression can be readily explained by differences in venous efflux anatomy. Together, our data are consistent with a model that edema formation is causally related to reductions in flow in the large draining veins, which in turn are caused by the accumulation of iRBCs.

Given the high sensitivity and prompt reactivity of the intracranial pressure to venous efflux obstructions³⁸ it seems possible that iRBC accumulation is a major factor leading to edema and brain death, particularly in cases with high parasite load. In infected patients, correlations were observed between brain swelling and parasite molecular markers in blood plasma related to iRBC biomass³⁹, while peripheral plasma cytokine levels did not correlate with brain swelling⁴⁰. Intriguingly, post-mortem studies have identified a subgroup of patients in which iRBC sequestration was the only pathological finding⁹. We hypothesize that iRBC accumulation in large draining veins and venous efflux obstructions are unifying hallmarks of onset of pediatric CM and ECM in murine models. Recent findings of increased hemoglobin concentrations, most likely representing vascular congestion in the microvasculature, as measured with near-infrared spectroscopy in infected children in vivo⁴¹, support this assumption and could represent a direct consequence of downstream obstructions.

Thus, inflammation accompanied by cytokine and chemokine production and CD8⁺ T cell infiltration are downstream events serving as robust pathological read-outs in ECM studies, but not necessarily being critical disease signatures in pediatric CM. Most importantly, recognition of reduced venous efflux in conjunction with iRBC sequestration lend strong support to the notion that murine malaria models can mimic CM and further emphasize the need to study disease onset prior to apparent clinical symptoms.

The effects of impairments in venous efflux in large draining veins are well known from cerebral venous thrombosis and mechanical obstructions of sinuses in both humans^38,42 and animal models^43,44,45. Hydrostatic pressure rises in upstream venous and capillary vessels and - depending on the presence or absence of collaterals - disruption of the blood-brain barrier, decreased perfusion, edema, and hemorrhages may follow.

Our data do not reveal whether the reduced efflux in the large draining veins was accompanied by thrombosis. In adult humans, sinus thrombosis in CM has been reported^46,47,48, but this is generally regarded a rare event⁴⁶. In African children, sinus thrombosis has neither been reported in post-mortem¹¹ nor in MRI studies⁴. The apparent lack of sinus thrombosis, however, does not preclude impairments of blood flow. This could be caused by combinations of iRBC sequestration, rosetting, and reduced deformability, which together likely impair fluid dynamics and lead to rheopathological effects that have been discussed earlier as major factors in CM pathology in both mice and humans^49,50. An MRI-study in Malawi children explicitly mentioned that transient occlusion of the internal cerebral vein and the vein of Galen could explain the pathology of a subgroup of patients⁴. We note that sequestration alone might not be sufficient to reduce the flow in large vessels to critical levels for inducing an edema. Additional alterations in viscosity and fluid dynamics are likely required, and the differences in flow observed between PbA-infected C57BL/6 wt and prf⁻^/⁻ mice might be attributed to cofactors, such as rosetting and hemozoin load.

There is broad consensus that the pathophysiology of CM is complex, and there is no one single disease signature in human post-mortem material or in neuroimaging in humans¹⁵. Thus far, studies and theories on CM pathology have focused on the microcirculation. Intravascular sequestration is the prime mechanism for iRBCs to avoid clearance by the spleen⁵¹, and of all vascular segments the capillary networks provide the largest surface area for sequestration. iRBC sequestration and/or rosetting in both the capillary and the venous space might be mediated by distinct or shared parasite-encoded proteins on the surface of infected erythrocytes. For instance, a member of the Plasmodium falciparum erythrocyte membrane protein 1 (PfEMP1) family that binds to endothelial protein C receptor (EPCR) has been linked to brain swelling in pediatric cerebral malaria³⁹, but potential candidates in murine malaria remain enigmatic. Clearly, future studies on the molecular mechanisms and parasite-host interactions during iRBC sequestration in large draining cerebral veins are needed. There is precedence in the human retina, where iRBC sequestration in large veins has been clearly demonstrated⁵². Several studies have shown that malaria retinopathies can indicate cerebral malaria in infected neurologically symptomatic patients^53,54, and it will be interesting to explore whether iRBC sequestration in large veins is a contributing factor in addition to occlusion of the retinal microvasculature and hypoxic injury. Of note, a similar sequestration phenotype has also been described in larger pial and subarachnoid vessels in human post-mortem material¹⁰. These observations together with our findings in the PbA-infected C57BL/6 wt mice suggest that a systematic analysis of the iRBC content in large veins in human post-mortem brains is warranted.

Edema in pediatric CM, as detected with MRI, is typically bilaterally symmetrical, affecting larger contiguous areas that do not correspond to arterial territories. It can be preferentially cortical with anterior or posterior phenotypes, subcortical or global^4,5,55,56. These diverse phenotypes can now be readily explained by venous efflux reductions in large draining veins with some individual variations in onset sites of critically reduced flow, i.e., superficial veins including bridging veins, sinuses, deep veins or globally affected large veins, rather than by varying predilection sites within the host microcirculation.

Especially in case of limited collateral flow, venous watershed areas could be particularly prone to damage, a phenomenon highlighted in human retinal pathology⁵⁷. It is of particular interest that the hemorrhages in children who died of CM are found in a confined subcortical white matter zone in post-mortem brains⁹. This subcortical white matter zone, which is also a well-delineated zone of corresponding MR-pathology in vivo⁵⁵, is a prominent venous watershed area between deep and superficial veins, an area also known for the so-called juxtacortical hemorrhages in cerebral venous thrombosis (see Fig. 4 in Coutinho et al. ⁵⁸).

In the murine malaria model, important events associated with ECM pathogenesis are intracerebral pro-inflammatory cytokine responses and influx of pathogenic CD8⁺ T cells^59,60,61. In good agreement, our study shows a progressive increase in cytokine levels starting at day 5 p.i. and peaking at day 7 p.i. when the mice succumb to the disease. The increase in cytokine levels was accompanied by an increase in a pro-inflammatory response, characterized by activation of brain parenchymal cells and infiltration of disease-causing CD8⁺ T cells, ultimately resulting in the disruption of the BBB. Anti-malarial treatment at day 5 p.i., when the peripheral parasitemia was already established, was sufficient to dampen the pro-inflammatory responses, indicating that accumulation of iRBC was essential for intracerebral pro-inflammatory response. To what extend cytokines occur in pediatric CM is not firmly established, largely due to the limitation that the measurements can only be performed in post-mortem tissue.

Notably, we detected iRBCs in the skull bone marrow. There is now consensus that bone marrow sequestration by iRBCs is a major site for parasite accumulation, gametocyte maturation, and, perhaps, parasite persistence in Plasmodium falciparum and Plasmodium vivax infections^62,63,64. Accordingly, iRBC accumulation in the skull bone marrow is expected, and we provide the first experimental evidence in a murine ECM model.

Our combined SPECT/CT and MR-angiography approach led to the identification of a previously unrecognized venous anatomy of the rostral superior sagittal sinus region, i.e., two interconnected parallel venous streams. Images in a number of studies from recent years indicate that this region is one of the nodal points of the lymphatic⁶⁵ and glymphatic⁶⁶ drainage systems and involved in bone marrow–brain interactions^67,68. The venous system in this area might, thus, share functional similarities not only to the superior sagittal sinus proper in humans but also to human perisinus spaces, e.g., venous lacunae, which are connected to diploic veins, as well as arachnoid granulations, in which recently lymphatic conduits and communication routes to the bone marrow have been demonstrated⁶⁹. These lacunae in humans are relatively large venous spaces that might be well-suited for iRBC sequestration or passive “sedimentation”. This could impair venous efflux through the bridging veins and might result in severe edema even with partially intact flow within the core of the superior sagittal sinus.

In summary, we show early stage iRBC sequestration and reduced flow in large draining veins only in P. berghei-infected brains that are on a trajectory to ECM. We propose that in CM and ECM pathology edema and hemorrhage formation are causally linked to iRBC-induced reductions in venous efflux in large draining veins downstream of the microcirculation and postcapillary venules.

Methods

Ethics statement

All animal experiments were in compliance with the German Animal Welfare Act (TierSchG) in a protocol approved by the state authorities (Landesverwaltungsamt Sachsen-Anhalt) file no: 42502-2-1440 UniMD.

Animals

Three different mouse strains were used in this study, C57BL/6 wt mice obtained from Janvier (Le Genest Saint Isle, France) and Jax, perforin-deficient C57BL/6 mice (C57BL/6-Prf1tm1Sdz/J, https://www.jax.org/strain/002407) and BALB/c mice (https://www.jax.org/strain/000651) obtained from Jax. All animals were kept in isolation under pathogen-free conditions in groups of 4 in standardized cages with 12 h of light-dark cycle and food and water was provided ad libitum in the animal facility of the Otto-von-Guericke University, Magdeburg, and the Leibniz Institute for Neurobiology, Magdeburg. For SPECT-imaging, mice were single-housed after implantation of the jugular vein catheter.

Parasite infection

The reference line ‘cl15cy1’ of the ANKA strain of P. berghei was used for the infection⁷⁰. For blood-stage infection, parasites were passaged in C57BL/6 mice (n = 20). At day 7 p.i., blood was collected through cardiac puncture, and blood stabilates were prepared by mixing 100 µl blood containing of 3 × 10⁶ iRBCs with 200 µl Alsever’s solution (Sigma–Aldrich, United Kingdom) and 10% glycerol (Calbiochem, Germany). For infection, 1 × 10⁶ PbA- iRBC (100 µl of blood mixed with Alsever’s solution) were injected intraperitoneally (i.p.) into each mouse. The infected mice were monitored daily from day 3 p.i. for clinical and neurological symptoms. The mice were evaluated based on the Rapid Murine Coma and Behaviour Scale (RMCBS)⁷¹. Mice were scored and divided into different groups based on their RMCBS: RMCBS score 15–20 (day 0–4 p.i.; asymptomatic), RMCBS score 10–14 (day 5–6 p.i.; early ECM) and RMCBS score 0–9 (day 7–8 p.i.; severe ECM). Parasitemia was measured daily after infection by microscopic examination of Diff-Quik (Medion Grifols Diagnostics, Switzerland) -stained thin blood smears.

Anti-malarial treatment

Starting at day 5 p.i., one group of PbA-infected mice was treated with the anti-malarial drug pyrimethamine (Sigma–Aldrich, Germany). 25 mg of pyrimethamine was dissolved in 1 ml of dimethyl sulfoxide (DMSO; Sigma–Aldrich, United Kingdom). 50 μl of this solution was dissolved with 950 μl of PBS (Gibco, United Kingdom) to make a working concentration of 1.25 mg/ml. Pyrimethamine was administered intravenously (i.v.) into mice (0.125 mg in 100 µl/mouse). The other group received phosphate-buffered saline (PBS) as control and has been referred to as untreated mice.

Evans blue staining

Mice were i.v. injected with 100 µl of 2% Evans blue solution (Sigma–Aldrich, USA) prepared in PBS (n = 2 per group of mice). One hour later, the mice were anesthetized with 4.5–5.0% isoflurane in 2:1 O₂:N₂O volume ratio and perfused transcardially with 0.9% NaCl (Carl Roth, Germany). The perfused brains were removed and photographed using a Sony Alpha A200K digital camera.

Quantitative real-time-PCR (q-PCR)

mRNA was isolated from the brains of the indicated groups of mice using an RNAeasy kit (Qiagen, Germany) (n = 4 per group). The SuperScript reverse transcriptase kit with oligo (dT) primers (Invitrogen, Canada) was used to transcribe the mRNA into cDNA. Quantitative-PCR for interferon-γ (IFN-γ), tumor necrosis factor (TNF), chemokine (C-X-C motif) ligand 3 (CXCL3), CXCL9, CXCL10, CXCL11, lymphotoxin α (LTα), LTβ and hypoxanthine phosphoribosyltransferase (HPRT) was performed using cDNA from olfactory bulb, cortex, and brainstem regions, respectively, of uninfected (day 0) and infected mice (day 5 and 7 p.i.) with and without pyrimethamine-treated mice employing the respective Taqman gene expression assay (Applied Biosystems, Germany). To determine the parasite load in the different regions of the brain, q-PCR was performed for P. berghei ANKA cytochrome B (PBANKA_MIT01900)⁷² and HPRT (Eurofins Genomics, Germany) using Light cycler 480 SyBr Green I master mix (Roche, Germany). Amplification was performed with a Light cycler 480 (Roche, Germany). Quantitation was performed using the Light cycler software SDS (version 5.1; Applied Biosystems, Germany), according to the ddC_T threshold cycle method with HPRT as the housekeeping gene⁷³. Data are depicted as the increase in the level of mRNA expression in infected mice over uninfected control mice.

SPECT/CT imaging

For SPECT-imaging, all animals were implanted, under isoflurane anesthesia (1.5–3% isoflurane, 850 ml/min O₂), with catheters in the right external jugular vein⁷⁴ and were given at least one day to recover.

SPECT-imaging of the biodistribution of ^99m Tc-labeled iRBCs

Blood was collected from C57BL/6 PbA-infected mice at day 7 p.i. with a parasite load of around 5% by cardiac puncture under isoflurane anesthesia. Enrichment of infected erythrocytes was performed by a Percoll gradient⁷⁵ with a yield of around 95% infected red blood cells. Around 7.5 × 10⁵ infected erythrocytes were incubated with freshly prepared ^99mTc-HMPAO for 30 min. Solutions were then centrifuged for 5 min at 500 g, the supernatant removed, and the cells resuspended in 0.9% NaCl. Cell viability was controlled using trypan blue staining. Awake mice were i.v. injected over 6 min with the labeled iRBCs (24.1 MBq on average) in volumes of 300 µl using perfusion pumps. Dynamic whole-body scans of 15 min duration starting shortly after injection were used in pilot experiments to determine spatial redistributions after injection. As no evidence for redistribution was found, mice (C57BL/6 wt d0 n = 7, 3 female, 4 male, mean age 11 weeks; d5 p.i., n = 8, 3 female, 5 male mean age 10.6 weeks; C57BL/6 prf⁻^/⁻ d0 n = 6, male, mean age 16.9 weeks, d5 p.i n = 6, male, mean age 20.2 weeks; BALB/c d0 n = 6, male, mean age 9.5 weeks, d5 p.i n = 6, male, mean age 9.5 weeks) were imaged with a single whole-body scan with an acquisition time of 20 min followed by a single head scan with an acquisition time of 90 min. After injection of the labeled iRBCs mice were anesthetized and transferred to the scanner. Scanning started with whole-body SPECT/CT followed by head SPECT/CT. The head SPECT started one hour after injection of the labeled iRBCs.

SPECT/CT imaging was performed with a four-head NanoSPECT/CT^TM scanner (Mediso, Hungary). Animals were scanned under gas anesthesia (1.2–1.5% isoflurane, 850 ml/min O₂). CT and SPECT were co-registered. Head scans were accompanied by two co-registered CT scans, one before and one after the SPECT scan, to control for motion artefacts (none detected). CT scans were made at 45 kVp, 177 μA, with 180 projections, 500 ms per projection, and 96 μm isotropic spatial resolution, reconstructed with the manufacturer´s software (InVivoScope 1.43) at isotropic voxel-sizes of 200 μm for whole-body scans and 100 µm for head scans. SPECT scans were made using nine-pinhole mouse brain apertures with 1.2 mm pinhole diameters providing a nominal spatial resolution ≤1 mm. Photopeaks were set to the default values of the NanoSPECT/CT for ^99mTc (140 keV ± 5%). SPECT images were reconstructed using the manufacturer´s software (HiSPECT^TM, SCIVIS, Germany) at an isotropic voxel size of 448 μm in whole-body scans and 300 µm in head scans.

For analysis, whole-body CTs were used to create, in the OsiriX^TM software, whole-body volume-of-interests (VOIs) for calculation of whole-body volumes and of ^99mTc-contents in the whole-body SPECT data. Whole-body ^99mTc-concentrations were calculated for each individual. All head SPECT data sets were aligned to a T2w anatomical reference MR. The alignment was based solely on the co-registered CT images. An automated workflow was used for the CT / MR alignment (see supplemental methods CT/SPECT registration). Briefly, the hyperintense skull was automatically segmented on CT images and the volume inside the skull was used to define a brain mask which was registered to the brain mask of the MRI. The calculated co-registration was then applied to the SPECT images.

For better analysis of the spatial relationship to the rostral rhinal vein all data sets were then aligned, using the same coordinate transformation, to a high-resolution MR dataset from the literature, the DSURQE-MR⁷⁶. This dataset is from an ex vivo contrast-agent perfused mouse, in which the rrv is clearly visible. SPECT head data sets were then divided by the according whole-body concentrations to get data sets in units of standardized uptake values (SUV). SUV in whole-brain VOIs and VOIs on the confluence of the sinuses and rrv were measured in OsiriX^TM (version 5.9.1, Pixmeo, Switzerland). Statistical testing using student t-test (unpaired, two-tailed, heteroscedastic) was done in Microsoft Excel^TM. Individual SPECT and CT head data sets of each group were added and divided by the number of animals per group in order to get group-mean data sets. Group-mean data sets and differences between group-mean SPECT data were overlaid on the anatomical reference MR and a group-mean MR-angiography from d0 mice in OsiriX^TM. Images from selected sections were arranged for illustrations in Photoshop^TM (version 24, Adobe, USA). For clarity, extracranial parts of the SPECT data were removed in the images.

SPECT-imaging of cerebral blood flow

Awake mice (d0 n = 10, male, mean age 10.5 weeks; d5 p.i. n = 9, male, mean age 9.8 weeks) during ongoing behavior were i.v. injected via the external jugular vein catheter with freshly prepared ^99mTc-HMPAO^74,77. Doses of on average 102 MBq were injected in 330 µl within 10 min using a perfusion pump.

SPECT/CT head scans and alignment of the SPECT data to the anatomical reference brain were made as described above with an acquisition time of 1 h. After alignment, SPECT data were converted to TIFF-files, brain data were cut out from the head scans using a brain mask and global-mean normalized in ImageJ. An unpaired two-tailed heteroscedastic voxel-wise t-test was made in MATLAB (version R2017b (9.3.0.713579) 64-bit, Mathworks, USA). T-test result files (uncorrected for multiple comparisons) as well as group-mean and difference-files were converted to DICOM and overlaid on the anatomical reference MR in OsiriX^TM. Specific sections were chosen and the figures were edited and arranged in Photoshop^TM.

MRI

Infected mice and controls underwent MR-imaging sessions including MR-TOF angiographies combined with T2w anatomical reference scans. Mice were scanned before infection, at day 5 p.i and at day 7 p.i. (C57BL/6 wt d0 n = 5, male, mean age 11 weeks, d5 n = 10, male, mean age 12.7 weeks, d7 n = 10, male, mean age 12.7 weeks; C57BL/6 prf⁻^/⁻ n = 6 male, mean age 15.8 weeks in each group; BALB/c n = 5, male, mean age 11 weeks in each group). Imaging of the brain was performed using a combination of a ¹H volume coil of 86 mm in inner diameter for transmission and a ¹H 3 × 1 surface array coil for receiving the signal. The MRI protocol included a MAPshim method for local shimming of the entire brain, a T2-weighted 2D TurboRARE (TR/TE = 4200 ms/18.5 ms, 2 averages) with equal field of view (25.6 × 25.6 mm) serving as anatomical reference for the equivalent MR-angiography. A non-contrast-enhanced 2D TOF with flow-compensation was acquired with optimized parameters (TR/TE: 3 ms/12 ms, in 80 slices with a slice thickness of 0.3 mm, slice overlaps of 0.1 mm and an in-plane-resolution of 0.08 × 0.08 mm resulting a scan time of ~4 min). The flip angle was set to 70° and the saturation slice, with a thickness of 3 mm, was positioned with a 1 mm gap towards the caudal side of the brain in order to image flow in caudo-rostral direction.

Anesthesia of the animals was induced with 3% Isoflurane carried by a 1:1 mixture of O₂/N₂O followed by a reduction to 1% Isoflurane while MR-imaging was performed. Vital parameters of the mice, such as body temperature and breathing rate, were permanently monitored and kept constant during scanning procedure.

For analysis, T2w and TOF images were aligned to the same anatomical MR reference space (see supplemental methods TOF-image registration). Briefly, a group average TOF was registered to each T2 MRI and used as a reference to align individual TOF images with corresponding T2-MR-images. T2-MR-images were then registered to the Allen brain atlas, the co-registration was then applied to the TOF to yield an angiogram in reference space.

Average group angiography images were made in the same manner as described in the SPECT/CT experiments. The files were overlaid in OsiriX^TM on MR T2 images of single individuals from each group. Images for the figures were arranged in Photoshop^TM (version 24).

For calculating the signal intensities in the superior sagittal sinus (sss), a volume-of interest (VOI) file was created manually in OsiriX^TM from all overlaid and aligned SSS in the control group. Mean signal intensities in the VOI in each individual were calculated using OsiriX^TM.

Histology

The histological analysis of the large draining veins was done on 7 µm thick Giemsa-stained paraffin sections from material fixed by immersion in 4% paraformaldehyde in 100 mM phosphate buffer (PB) pH 7.4 and decalcified in Surgipath^TM Decalcifier II (Leica, Germany). Mice were euthanized with isoflurane. The skin overlaying the dorsal cranium was removed. For better penetration of the fixative parts of the cranium were removed in parietal regions. In addition, a small whole of about 1 mm diameter was drilled into the frontal bones on each side. Care was taken to leave the bone intact above sinuses and the rostral rhinal vein. When removing bones, a distance of at least 1 mm to the sinuses was kept. Material from non-exsanguinated animals was used. After skull preparation the entire bodies were fixed in 4% PFA. The heads were removed from the bodies after fixation and the remaining skin was removed. All heads were stored in 1% PFA for no longer than five days. Heads were washed 3 × 10 min in PB, decalcified for 2 days, washed 3 × 10 min in 0.9% NaCl, dehydrated in an ascending ethanol series, and embedded in paraffin (Paraplast Plus, Roth, Germany). Seven micrometer thick whole-head sections were made on a sliding microtome (HM 400). Sections were mounted on gelatinized Superfrost® Plus glass slides that were covered with a drop of an albumin-glycerol solution before mounting. After drying sections were Giemsa-stained (Giemsa-solution, Roth, Germany) and covered with Merckoglas® and coverslips. Data from PbA-infected C57BL/6 wt mice (n = 3 at d5, 2 female, 1 male, mean age 9 weeks; n = 1 at d6 p.i. female, 9 weeks; n = 3 at d7 p.i. 2 female, 1 male, mean age 9 weeks), C57BL/6 prf⁻^/⁻ mice (n = 3 at d7 p.i.,male, mean age 9.7 weeks) and PbNK65-infected C57BL/6 wt mice (n = 4 at d7 p.i., male, mean age 8 weeks) were analyzed microscopically. Photomicrographs were taken with a Histoscanner (3DHistech). High-resolution oil immersion images with and without polarizing filters were made on a Leica DMR microscope. The findings as displayed in Figs. 4, 5, and S2 were reproducible in each animal in the according groups. The anatomical relationships displayed in Fig. 6 were present in all mice studied in these groups.

Immunohistochemistry (IHC)

For immunohistochemistry, mice were perfused transcardially with 0.9% NaCl followed by 4% paraformaldehyde (PFA; Carl Roth, Germany) solution in PBS (n = 3 per group and time point) after isoflurane anesthesia (4.5–5.0% isoflurane in 2:1 O₂:N₂O volume ratio). The brains were removed, post-fixed in 4% PFA solution for 48 h, and cryoprotected in 30% sucrose (Sigma–Aldrich, USA) solution for 72 h followed by freezing in isopentane cooled by liquid nitrogen. Sagittal sections (50 µm) of frozen brain tissue were cut on a microtome (Leica CM 1950, Leica, Germany) and processed for hematoxylin (Mayer’s Hematoxylin solution, Sigma–Aldrich, USA) and eosin (Merck, Germany) (H&E) staining. Immunostaining was performed with primary antibodies against glial fibrillary acidic protein (GFAP; rabbit, 1:10000, #z0334, Dako Denmark AS, Denmark), ionized calcium-binding adapter molecule 1 (Iba1; rabbit, 1:2000, #019-19741, Wako Chemicals GmbH, Germany), cleaved caspase-3 (Casp3; rabbit, 1:500, #9661 L, Cell Signalling Technology Europe B.V., Germany), CD31 (rat, 1:2000, #550274, BD Biosciences, Germany), and CD8 (rat, 1:1000, #14-0808-82, eBioscience, Affymetrix Inc., USA), followed by staining with the appropriate biotinylated secondary antibodies (anti-rabbit: 1:200, #111-065-144, Dianova, Germany and anti-rat: 1:200, #712-065-153, Dianova, Germany), and avidin-biotin-peroxidase complex (ABC kit, Vector Laboratories, USA). Diaminobenzidine (DAB; Sigma–Aldrich, USA) was used to visualize the antibody reactions. The sections were mounted on gelatine-coated slides and covered using coverslips and mounting solution, Merckoglas (Merck, Germany). Images of different brain regions including olfactory bulb (OB), sub-ventricular zone (SVZ), rostral migratory stream (RMS), hippocampus, midbrain and pons, were acquired using a digital camera (Leica DFC 500, Leica, Germany) mounted on a combined brightfield and epifluorescence microscope (Zeiss Axioskop 2, Zeiss, Germany).

For bromodeoxyuridine and doublecortin immunofluorescence (IF) staining bromodeoxyuridine (BrdU; 50 μg/g body weight; eBioscience, USA) was i.p. administered at day 3 and day 5 p.i. The mice were transcardially perfused 48 h post injection, (n = 3 per group), as aforementioned, with 0.9% NaCl and 4% PFA after isoflurane anesthesia. Brains were fixed in 4% PFA for 48 h followed by cryoprotection in 30% sucrose solution for 72 h and were frozen in liquid nitrogen-cooled isopentane. Sagittal sections (50 μm) were immunostained with primary antibodies against doublecortin (DCX; goat, 1:1000, #sc-8066, Santa Cruz Biotechnology, USA) followed by incubation with immunofluorescent Alexa-488 labeled secondary antibody (anti-goat, 1:200, #A11055, Invitrogen by Thermo Fischer Scientific, USA). The sections were subsequently incubated in 2 N HCl at 37 °C for 1 h to denature the genomic DNA followed by neutralization in 0.1 M borate buffer. Primary antibodies against BrdU (rat, 1:2000, #ab6326, Abcam, United Kingdom) were added and counterstained by immunofluorescent Cy3 labeled secondary antibody (anti-rat, 1:200, #712-165-153, Dianova, Germany). Sections were placed on gelatin-coated slides and covered with coverslips and mounting solution, MOWIOL (Fluka, Germany). The rostral migratory stream (RMS) was imaged with a digital camera (Leica DFC 500, Leica, Germany) mounted on the combined brightfield and epifluorescence microscope (Zeiss Axioskop 2, Zeiss, Germany).

For IHC analysis, every single sagittal brain section was photographed with a digital camera (Leica DFC 500, Leica, Germany) mounted on a combined brightfield and epifluorescence microscope (Zeiss Axioskop 2, Zeiss, Germany). For H&E staining, images were acquired using 20× objective under brightfield illumination with exposure time of 8 ms. Cy3 conjugated BrdU antibodies were excited at 570 nm and Alexa-488 conjugated DCX antibodies were excited at 493 nm and images were captured using 5× objective in dark with exposure time of 590 ms and 650 ms, respectively. Images were acquired using a 20× objective under constant brightfield illumination for GFAP, Iba1, CD31, CD8 and cleaved Caspase-3 staining and their exposure times were 10 ms, 8 ms, 7 ms, 8 ms and 7 ms respectively. The respective images acquired were used for cell counting. Magnified images were captured under 40× objective for all the staining. For analysis and cell counting, ImageJ software (NIH, USA) was employed. The RGB images were converted to 16-bit gray scale images and threshold values were adjusted for each individual stain to highlight the structures to be counted. To reduce background noise in the images, background subtraction with rolling ball was carried out and particles were analyzed. With the exception of CD8 staining, cell number for all other staining were measured by automated counting the area covered by each cell type in the whole area of the image. For CD8 staining, the total number of CD8⁺ cells in the whole image were counted manually with the Cell Counter plugin in ImageJ.

CT/SPECT registration

The image registration to standard space (Allen Mouse Brain atlas 2017 (CCFv3); http://help.brain-map.org/display/mouseconnectivity/API) was performed using the ANTx2 toolbox (https://github.com/ChariteExpMri/antx2) running in MATLAB (R2016b, Natick, Massachusetts, The MathWorks Inc.). Detailed information for registration can be found in Koch et al. ²⁸. For image registration to standard space (Fig. S12), the T2w image was rigidly registered to the Allen Brain template and segmented into the three tissue compartments (gray matter, white matter, and cerebrospinal fluid) using SPM’s unified segmentation approach⁷⁸. The tissue segments where further used for affine and subsequent nonlinear registration to the template tissue probability maps⁷⁹ using elastix⁸⁰ to obtain the T2w image in standard space.

The inverse transformation parameters based on the transformation parameters from rigid, affine, and nonlinear forward registration was used to back-transform the binary mask of the Allen mouse brain to the animal native space. The skull from the animal’s CT image was extracted using Otsu and watershed segmentations, followed by image clustering and morphological image operations to finally obtain a binary CT-based brain mask. In the next step, the CT image and the SPECT image (in register with the CT image) were brought into to the animal’s native space (T2w space). For this, the CT-based brain mask was rigidly registered to the Allen brain mask in native space and the same transformation parameters were the applied to the SPECT image. Finally, CT and SPECT images in native space were transformed to standard space by applying the forward transform parameters obtained from the first processing step. The CT/SPECT image registration workflow is implemented in the ANTx2 toolbox.

TOF-image registration

The image registration to standard space (Allen Mouse Brain atlas 2017 (CCFv3); http://help.brain-map.org/display/mouseconnectivity/API) was performed using the ANTx2 toolbox (https://github.com/ChariteExpMri/antx2) running in MATLAB (R2016b, Natick, Massachusetts, The MathWorks Inc.). Detailed information for registration can be found in Koch et al. ²⁸. We first generated a TOF template by manual registration of TOF images from a group of animals to an individual T2w image using the MPI-Tool^TM software (version 6.36, ATV, Advanced Tomo Vision, D-50169 Kerpen, Germany). For registration of the TOF template to standard space, the T2-weighted (T2w) image was rigidly registered to the Allen brain template, and segmented into the three tissue compartments (gray matter, white matter, and cerebrospinal fluid) using SPM’s (https://www.fil.ion.ucl.ac.uk/spm/) unified segmentation approach⁷⁸. The tissue segments where further used for affine and subsequent nonlinear registration to the template tissue probability maps⁷⁹ using elastix⁷² to obtain the T2w image in standard space. Using the concatenated transformation parameters from rigid, affine, and nonlinear registrations, the TOF template was finally transformed to standard space. For each animal, the T2w image from native space was transformed to standard space as described above. In a next step, the TOF template in standard space was back-transformed to each animal’s native space using the inverse transformation parameters for registration to standard space. Next, the animal’s TOF-image was rigidly registered to the TOF template in the animal’s native space. Finally, the registered TOF-image was transformed from native space to standard space using the animal’s concatenated transformation parameters (rigid, affine, and nonlinear registrations).

For each animal, the T2w image from native space was transformed to standard space as described above. In a next step, the TOF template in standard space was back-transformed to each animal’s native space using the inverse transformation parameters for registration to standard space. Next, the animal’s TOF-image was rigidly registered to the TOF template in the animal’s native space. Finally, the registered TOF-image was transformed from native space to standard space using the animal’s concatenated transformation parameters (rigid, affine, and nonlinear registrations).

Statistics

Statistical analyses of percentage of appearance of symptoms, parasitemia, immunohistochemistry, and q-PCR, respectively, were performed using GraphPad Prism versions 9 and 10 (GraphPad, USA). Statistical differences between groups were determined using two-way ANOVA. All experiments were performed at least twice. P-values < 0.05 were considered to be * significant, P-values < 0.01 were ** significant, and P-values < 0.001 were *** significant.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

All primary data associated with this study are available in the article and the supplementary information. The source data are provided as a Source Data file at: https://doi.org/10.6084/m9.figshare.25225802

References

World Health Organization. World Malaria Report 2023 (World Heath Organization, 2023).
Brewster, D. R., Kwiatkowski, D. & White, N. J. Neurological sequelae of cerebral malaria in children. Lancet 336, 1039–1043 (1990).
Article CAS PubMed Google Scholar
Idro, R., Jenkins, N. E. & Newton, C. R. Pathogenesis, clinical features, and neurological outcome of cerebral malaria. Lancet Neurol. 4, 827–840 (2005).
Article PubMed Google Scholar
Seydel, K. B. et al. Brain swelling and death in children with cerebral malaria. N. Engl. J. Med. 372, 1126–1137 (2015).
Article CAS PubMed PubMed Central Google Scholar
Potchen, M. J. et al. Acute brain MRI findings in 120 Malawian children with cerebral malaria: new insights into an ancient disease. AJNR Am. J. Neuroradiol. 33, 1740–1746 (2012).
Article CAS PubMed PubMed Central Google Scholar
Kampondeni, S. D. et al. Noninvasive measures of brain edema predict outcome in pediatric cerebral malaria. Surg. Neurol. Int. 9, 53 (2018).
Article PubMed PubMed Central Google Scholar
MacPherson, G. G., Warrell, M. J., White, N. J., Looareesuwan, S. & Warrell, D. A. Human cerebral malaria. A quantitative ultrastructural analysis of parasitized erythrocyte sequestration. Am. J. Pathol. 119, 385–401 (1985).
CAS PubMed PubMed Central Google Scholar
Silamut, K. et al. A quantitative analysis of the microvascular sequestration of malaria parasites in the human brain. Am. J. Pathol. 155, 395–410 (1999).
Article CAS PubMed PubMed Central Google Scholar
Taylor, T. E. et al. Differentiating the pathologies of cerebral malaria by postmortem parasite counts. Nat. Med. 10, 143–145 (2004).
Article CAS PubMed Google Scholar
Dorovini-Zis, K. et al. The neuropathology of fatal cerebral malaria in Malawian children. Am. J. Pathol. 178, 2146–2158 (2011).
Article PubMed PubMed Central Google Scholar
Milner, D. A. Jr. et al. The systemic pathology of cerebral malaria in African children. Front. Cell. Infect. Microbiol. 4, 104 (2014).
Article PubMed PubMed Central ADS Google Scholar
Haldar, K., Murphy, S. C., Milner, D. A. & Taylor, T. E. Malaria: mechanisms of erythrocytic infection and pathological correlates of severe disease. Annu. Rev. Pathol. 2, 217–249 (2007).
Article CAS PubMed Google Scholar
Nishanth, G. & Schluter, D. Blood-brain barrier in cerebral malaria: pathogenesis and therapeutic intervention. Trends Parasitol. 35, 516–528 (2019).
Article CAS PubMed Google Scholar
Strangward, P. et al. A quantitative brain map of experimental cerebral malaria pathology. PLoS Pathog. 13, e1006267 (2017).
Article PubMed PubMed Central Google Scholar
Taylor, T. E. & Molyneux, M. E. The pathogenesis of pediatric cerebral malaria: eye exams, autopsies, and neuroimaging. Ann. N. Y. Acad. Sci. 1342, 44–52 (2015).
Article CAS PubMed PubMed Central ADS Google Scholar
White, N. J., Turner, G. D., Medana, I. M., Dondorp, A. M. & Day, N. P. The murine cerebral malaria phenomenon. Trends Parasitol. 26, 11–15 (2010).
Article PubMed PubMed Central Google Scholar
Langhorne, J. et al. The relevance of non-human primate and rodent malaria models for humans. Malar. J. 10, 23 (2011).
Article PubMed PubMed Central Google Scholar
Craig, A. G. et al. The role of animal models for research on severe malaria. PLoS Pathog. 8, e1002401 (2012).
Article CAS PubMed PubMed Central Google Scholar
Amante, F. H. et al. Immune-mediated mechanisms of parasite tissue sequestration during experimental cerebral malaria. J. Immunol. 185, 3632–3642 (2010).
Article CAS PubMed Google Scholar
Claser, C. et al. CD8+ T cells and IFN-gamma mediate the time-dependent accumulation of infected red blood cells in deep organs during experimental cerebral malaria. PLoS ONE 6, e18720 (2011).
Article CAS PubMed PubMed Central ADS Google Scholar
Howland, S. W. et al. Brain microvessel cross-presentation is a hallmark of experimental cerebral malaria. EMBO Mol. Med. 5, 984–999 (2013).
Article PubMed Google Scholar
Khoury, D. S. et al. Effect of mature blood-stage Plasmodium parasite sequestration on pathogen biomass in mathematical and in vivo models of malaria. Infect. Immun. 82, 212–220 (2014).
Article PubMed PubMed Central Google Scholar
Matz, J. M. et al. The Plasmodium berghei translocon of exported proteins reveals spatiotemporal dynamics of tubular extensions. Sci. Rep. 5, 12532 (2015).
Article CAS PubMed PubMed Central ADS Google Scholar
Possemiers, H. et al. Skeleton binding protein-1-mediated parasite sequestration inhibits spontaneous resolution of malaria-associated acute respiratory distress syndrome. PLoS Pathog. 17, e1010114 (2021).
Article CAS PubMed PubMed Central Google Scholar
Dorr, A., Sled, J. G. & Kabani, N. Three-dimensional cerebral vasculature of the CBA mouse brain: a magnetic resonance imaging and micro computed tomography study. Neuroimage 35, 1409–1423 (2007).
Article CAS PubMed Google Scholar
Nitcheu, J. et al. Perforin-dependent brain-infiltrating cytotoxic CD8+ T lymphocytes mediate experimental cerebral malaria pathogenesis. J. Immunol. 170, 2221–2228 (2003).
Article CAS PubMed Google Scholar
Cimperman, C. K. et al. Cerebral malaria is regulated by host-mediated changes in plasmodium gene expression. mBio 14, e0339122 (2023).
Article PubMed Google Scholar
Koch, S. et al. Atlas registration for edema-corrected MRI lesion volume in mouse stroke models. J. Cereb. Blood Flow. Metab. 39, 313–323 (2019).
Article PubMed Google Scholar
Hoffmann, A. et al. Experimental cerebral malaria spreads along the rostral migratory stream. PLoS Pathog. 12, e1005470 (2016).
Article PubMed PubMed Central Google Scholar
Huggins, M. A. et al. Perforin expression by CD8 T cells is sufficient to cause fatal brain edema during experimental cerebral malaria. Infect. Immun. 85, e00985–16 (2017).
Article CAS PubMed PubMed Central Google Scholar
De Niz, M. et al. Plasmodium gametocytes display homing and vascular transmigration in the host bone marrow. Sci. Adv. 4, eaat3775 (2018).
Article PubMed PubMed Central ADS Google Scholar
Xiong, B. et al. Precise cerebral vascular atlas in stereotaxic coordinates of whole mouse brain. Front. Neuroanat. 11, 128 (2017).
Article PubMed PubMed Central Google Scholar
Mancini, M. et al. Head and neck veins of the mouse. A magnetic resonance, micro computed tomography and high frequency color Doppler ultrasound study. PLoS ONE 10, e0129912 (2015).
Article PubMed PubMed Central Google Scholar
Nie, C. Q. et al. IP-10-mediated T cell homing promotes cerebral inflammation over splenic immunity to malaria infection. PLoS Pathog. 5, e1000369 (2009).
Article PubMed PubMed Central Google Scholar
Adams, Y. et al. Plasmodium falciparum erythrocyte membrane protein 1 variants induce cell swelling and disrupt the blood-brain barrier in cerebral malaria. J. Exp. Med. 218, e20201266 (2021).
Article CAS PubMed PubMed Central Google Scholar
Lee, W. C., Russell, B. & Renia, L. Evolving perspectives on rosetting in malaria. Trends Parasitol. 38, 882–889 (2022).
Article CAS PubMed Google Scholar
Rodriguez-Munoz, D. et al. Hypothyroidism confers tolerance to cerebral malaria. Sci. Adv. 8, eabj7110 (2022).
Article CAS PubMed PubMed Central Google Scholar
Wilson, M. H. Monro-Kellie 2.0: The dynamic vascular and venous pathophysiological components of intracranial pressure. J. Cereb. Blood Flow. Metab. 36, 1338–1350 (2016).
Article PubMed PubMed Central Google Scholar
Kessler, A. et al. Linking EPCR-binding PfEMP1 to brain swelling in pediatric cerebral malaria. Cell Host Microbe 22, 601–614 e605 (2017).
Article CAS PubMed PubMed Central Google Scholar
Harawa, V. et al. Brain swelling is independent of peripheral plasma cytokine levels in Malawian children with cerebral malaria. Malar. J. 17, 435 (2018).
Article CAS PubMed PubMed Central Google Scholar
Smith, R. L. et al. Increased brain microvascular hemoglobin concentrations in children with cerebral malaria. Sci. Transl. Med. 15, eadh4293 (2023).
Article CAS PubMed Google Scholar
Silvis, S. M., de Sousa, D. A., Ferro, J. M. & Coutinho, J. M. Cerebral venous thrombosis. Nat. Rev. Neurol. 13, 555–565 (2017).
Article PubMed Google Scholar
Ungersbock, K., Heimann, A. & Kempski, O. Cerebral blood flow alterations in a rat model of cerebral sinus thrombosis. Stroke 24, 569–570 (1993).
Article Google Scholar
Gotoh, M., Ohmoto, T. & Kuyama, H. Experimental study of venous circulatory disturbance by dural sinus occlusion. Acta Neurochir. 124, 120–126 (1993).
Article CAS PubMed Google Scholar
Ueda, K., Nakase, H., Miyamoto, K., Otsuka, H. & Sakaki, T. Impact of anatomical difference of the cerebral venous system on microcirculation in a gerbil superior sagittal sinus occlusion model. Acta Neurochir. 142, 75–82 (2000).
Article CAS PubMed Google Scholar
Krishnan, A., Karnad, D. R., Limaye, U. & Siddharth, W. Cerebral venous and dural sinus thrombosis in severe falciparum malaria. J. Infect. 48, 86–90 (2004).
Article CAS PubMed Google Scholar
Luvira, V., Chamnanchanunt, S., Thanachartwet, V., Phumratanaprapin, W. & Viriyavejakul, A. Cerebral venous sinus thrombosis in severe malaria. Southeast Asian J. Trop. Med. Public Health 40, 893–897 (2009).
PubMed Google Scholar
Pinzon, M. A., Pineda, J. C., Rosso, F., Shinchi, M. & Bonilla-Abadia, F. Plasmodium vivax cerebral malaria complicated with venous sinus thrombosis in Colombia. Asian Pac. J. Trop. Med. 6, 413–415 (2013).
Article PubMed Google Scholar
Russell, B. M. & Cooke, B. M. The rheopathobiology of plasmodium vivax and other important primate malaria parasites. Trends Parasitol. 33, 321–334 (2017).
Article PubMed Google Scholar
Herricks, T., Seydel, K. B., Molyneux, M., Taylor, T. & Rathod, P. K. Estimating physical splenic filtration of Plasmodium falciparum-infected red blood cells in malaria patients. Cell. Microbiol. 14, 1880–1891 (2012).
Article CAS PubMed PubMed Central Google Scholar
Miller, L. H., Ackerman, H. C., Su, X. Z. & Wellems, T. E. Malaria biology and disease pathogenesis: insights for new treatments. Nat. Med. 19, 156–167 (2013).
Article CAS PubMed PubMed Central Google Scholar
Lewallen, S. et al. Clinical-histopathological correlation of the abnormal retinal vessels in cerebral malaria. Arch. Ophthalmol. 118, 924–928 (2000).
CAS PubMed Google Scholar
Lewallen, S. et al. Retinal findings predictive of outcome in cerebral malaria. Trans. R. Soc. Trop. Med. Hyg. 90, 144–146 (1996).
Article CAS PubMed Google Scholar
Beare, N. A., Harding, S. P., Taylor, T. E., Lewallen, S. & Molyneux, M. E. Perfusion abnormalities in children with cerebral malaria and malarial retinopathy. J. Infect. Dis. 199, 263–271 (2009).
Article PubMed Google Scholar
Potchen, M. J. et al. 1.5 tesla magnetic resonance imaging to investigate potential etiologies of brain swelling in pediatric cerebral malaria. Am. J. Trop. Med. Hyg. 98, 497–504 (2018).
Article CAS PubMed PubMed Central Google Scholar
Moghaddam, S. M. et al. Diffusion-weighted MR imaging in a prospective cohort of children with cerebral malaria offers insights into pathophysiology and prognosis. AJNR Am. J. Neuroradiol. 40, 1575–1580 (2019).
CAS PubMed PubMed Central Google Scholar
MacCormick, I. J. et al. Cerebral malaria in children: using the retina to study the brain. Brain 137, 2119–2142 (2014).
Article PubMed PubMed Central Google Scholar
Coutinho, J. M. et al. Small juxtacortical hemorrhages in cerebral venous thrombosis. Ann. Neurol. 75, 908–916 (2014).
Article PubMed Google Scholar
Belnoue, E. et al. On the pathogenic role of brain-sequestered alphabeta CD8+ T cells in experimental cerebral malaria. J. Immunol. 169, 6369–6375 (2002).
Article CAS PubMed Google Scholar
Renia, L. et al. Pathogenic T cells in cerebral malaria. Int. J. Parasitol. 36, 547–554 (2006).
Article CAS PubMed Google Scholar
Dunst, J., Kamena, F. & Matuschewski, K. Cytokines and chemokines in cerebral malaria pathogenesis. Front. Cell. Infect. Microbiol. 7, 324 (2017).
Article PubMed PubMed Central Google Scholar
Joice, R. et al. Plasmodium falciparum transmission stages accumulate in the human bone marrow. Sci. Transl. Med. 6, 244re245 (2014).
Article Google Scholar
Obaldia, N. 3rd et al. Bone marrow is a major parasite reservoir in plasmodium vivax infection. mBio 9, e00625–18 (2018).
Article CAS PubMed PubMed Central Google Scholar
Farfour, E., Charlotte, F., Settegrana, C., Miyara, M. & Buffet, P. The extravascular compartment of the bone marrow: a niche for Plasmodium falciparum gametocyte maturation? Malar. J. 11, 285 (2012).
Article PubMed PubMed Central Google Scholar
Jacob, L. et al. Conserved meningeal lymphatic drainage circuits in mice and humans. J. Exp. Med. 219, e20220035 (2022).
Article CAS PubMed PubMed Central Google Scholar
Hablitz, L. M. et al. Increased glymphatic influx is correlated with high EEG delta power and low heart rate in mice under anesthesia. Sci. Adv. 5, eaav5447 (2019).
Article CAS PubMed PubMed Central ADS Google Scholar
Herisson, F. et al. Direct vascular channels connect skull bone marrow and the brain surface enabling myeloid cell migration. Nat. Neurosci. 21, 1209–1217 (2018).
Article CAS PubMed PubMed Central Google Scholar
Mazzitelli, J. A. et al. Cerebrospinal fluid regulates skull bone marrow niches via direct access through dural channels. Nat. Neurosci. 25, 555–560 (2022).
Article CAS PubMed PubMed Central Google Scholar
Shah, T. et al. Arachnoid granulations are lymphatic conduits that communicate with bone marrow and dura-arachnoid stroma. J. Exp. Med. 220, e20220618 (2023).
Article CAS PubMed Google Scholar
Janse, C. J., Ramesar, J. & Waters, A. P. High-efficiency transfection and drug selection of genetically transformed blood stages of the rodent malaria parasite Plasmodium berghei. Nat. Protoc. 1, 346–356 (2006).
Article CAS PubMed Google Scholar
Carroll, R. W. et al. A rapid murine coma and behavior scale for quantitative assessment of murine cerebral malaria. PLoS ONE 5, e13124 (2010).
Article PubMed PubMed Central ADS Google Scholar
Bahamontes-Rosa, N. et al. New molecular settings to support in vivo anti-malarial assays. Malar. J. 15, 147 (2016).
Article PubMed PubMed Central Google Scholar
Livak, K. J. & Schmittgen, T. D. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-delta delta C(T)) method. Methods 25, 402–408 (2001).
Article CAS PubMed Google Scholar
Oelschlegel, A. M. & Goldschmidt, J. Functional neuroimaging in rodents using cerebral blood flow SPECT. Front. Phys. 8 152 (2020).
Trang, D. T., Huy, N. T., Kariu, T., Tajima, K. & Kamei, K. One-step concentration of malarial parasite-infected red blood cells and removal of contaminating white blood cells. Malar. J. 3, 7 (2004).
Article PubMed PubMed Central Google Scholar
Dorr, A. E., Lerch, J. P., Spring, S., Kabani, N. & Henkelman, R. M. High resolution three-dimensional brain atlas using an average magnetic resonance image of 40 adult C57Bl/6J mice. Neuroimage 42, 60–69 (2008).
Article CAS PubMed Google Scholar
Kolodziej, A. et al. SPECT-imaging of activity-dependent changes in regional cerebral blood flow induced by electrical and optogenetic self-stimulation in mice. Neuroimage 103, 171–180 (2014).
Article PubMed Google Scholar
Ashburner, J. & Friston, K. J. Unified segmentation. Neuroimage 26, 839–851 (2005).
Article PubMed Google Scholar
Hikishima, K. et al. In vivo microscopic voxel-based morphometry with a brain template to characterize strain-specific structures in the mouse brain. Sci. Rep. 7, 85 (2017).
Article PubMed PubMed Central ADS Google Scholar
Klein, S., Staring, M., Murphy, K., Viergever, M. A. & Pluim, J. P. elastix: a toolbox for intensity-based medical image registration. IEEE Trans. Med. Imaging 29, 196–205 (2010).
Article PubMed Google Scholar

Download references

Acknowledgements

This project has been funded by the federal state of Saxony-Anhalt, Germany, and the European Structural and Investment Funds (ESF, 2014-2020), project number ZS/2016/08/80645 to G.N., D.S., E.B., and J.G. and the Deutsche Forschungsgemeinschaft (SFB 854, TP30 to G.N.). Funding to S.P.K. and P.B.S. was provided by the German Federal Ministry of Education and Research (BMBF) under the ERA-NET NEURON scheme (01EW1811 and 01EW2305), and the German Research Foundation (DFG, Project BO 4484/2-1 and EXC-2049-390688087 NeuroCure). Analysis of imaging data was supported by Charité 3R | Replace, Reduce, Refine. The authors wish to thank Holger Reim, Janet Stallmann, Anja Gürke, Marko Dombach, Gordon Kühne, Torsten Stoeter (Leibniz Institute for Neurobiology, Magdeburg), Annette Sohnekind, Nadja Schlüter, Anita Marquardt (Institute for Medical Microbiology, University Hospital, Magdeburg) for their excellent support and technical assistance in the experiments.

Funding

Open Access funding enabled and organized by Projekt DEAL.

Author information

These authors contributed equally: A. M. Oelschlegel, R. Bhattacharjee, P. Wenk.

Authors and Affiliations

Combinatorial NeuroImaging Core Facility, Leibniz Institute for Neurobiology, 39118, Magdeburg, Germany
A. M. Oelschlegel, R. Bhattacharjee, P. Wenk, E. Budinger & J. Goldschmidt
Research group Neuroplasticity, Leibniz Institute for Neurobiology, 39118, Magdeburg, Germany
A. M. Oelschlegel
Institute of Medical Microbiology and Hospital Epidemiology, Hannover Medical School, 30625, Hannover, Germany
R. Bhattacharjee, K. Harit, D. Schlüter & G. Nishanth
Institute of Anatomy, Medical Faculty, Otto-von-Guericke-University Magdeburg, Leipziger Strasse 44, 39120, Magdeburg, Germany
H-J Rothkötter
Charité-Universitätsmedizin Berlin, corporate member of Freie Universität Berlin and Humboldt-Universität zu Berlin, Department of Experimental Neurology and Center for Stroke Research, Charitéplatz 1, 10117, Berlin, Germany
S. P. Koch & P. Boehm-Sturm
Charité-Universitätsmedizin Berlin, NeuroCure Cluster of Excellence and Charité Core Facility 7T Experimental MRIs, 10117, Berlin, Germany
S. P. Koch & P. Boehm-Sturm
Charité-Universitätsmedizin Berlin, Charité 3R | Replace, Reduce, Refine, Charitéplatz 1, 10117, Berlin, Germany
S. P. Koch & P. Boehm-Sturm
Department of Molecular Parasitology, Institute of Biology, Humboldt University, 10115, Berlin, Germany
K. Matuschewski
Center of Behavioural Brain Sciences, Universitätsplatz 2, 39106, Magdeburg, Germany
E. Budinger & J. Goldschmidt

Authors

A. M. Oelschlegel
View author publications
You can also search for this author in PubMed Google Scholar
R. Bhattacharjee
View author publications
You can also search for this author in PubMed Google Scholar
P. Wenk
View author publications
You can also search for this author in PubMed Google Scholar
K. Harit
View author publications
You can also search for this author in PubMed Google Scholar
H-J Rothkötter
View author publications
You can also search for this author in PubMed Google Scholar
S. P. Koch
View author publications
You can also search for this author in PubMed Google Scholar
P. Boehm-Sturm
View author publications
You can also search for this author in PubMed Google Scholar
K. Matuschewski
View author publications
You can also search for this author in PubMed Google Scholar
E. Budinger
View author publications
You can also search for this author in PubMed Google Scholar
D. Schlüter
View author publications
You can also search for this author in PubMed Google Scholar
J. Goldschmidt
View author publications
You can also search for this author in PubMed Google Scholar
G. Nishanth
View author publications
You can also search for this author in PubMed Google Scholar

Contributions

G.N. and J.G. conceptualized and supervised the experiments. A.M.O., R.B., P.W., K.H., H.R., S.K., P.B., J.G. and G.N. performed experiments and analyzed data. A.M.O., R.B., P.W., K.H., K.M., E.B., D.S., J.G. and G.N. interpreted the data. G.N. and J.G. wrote the manuscript. D.S. and K.M. reviewed and edited the manuscript.

Corresponding authors

Correspondence to J. Goldschmidt or G. Nishanth.

Ethics declarations

Competing interests

The authors declare no competing interests.

Peer review

Peer review information

Nature Communications thanks Aubrey Cunnington, Terrie Taylor, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Supplementary Information

Peer Review File

Reporting Summary

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.

Reprints and permissions

About this article

Cite this article

Oelschlegel, A.M., Bhattacharjee, R., Wenk, P. et al. Beyond the microcirculation: sequestration of infected red blood cells and reduced flow in large draining veins in experimental cerebral malaria. Nat Commun 15, 2396 (2024). https://doi.org/10.1038/s41467-024-46617-w

Download citation

Received: 24 February 2022
Accepted: 29 February 2024
Published: 16 March 2024
DOI: https://doi.org/10.1038/s41467-024-46617-w

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.