Here, we report the existence of meningeal lymphatic vessels in human and nonhuman primates (common marmoset monkeys) and the feasibility of noninvasively imaging and mapping them in vivo with high-resolution, clinical MRI. On T2-FLAIR and T1-weighted black-blood imaging, lymphatic vessels enhance with gadobutrol, a gadolinium-based contrast agent with high propensity to extravasate across a permeable capillary endothelial barrier, but not with gadofosveset, a blood-pool contrast agent. The topography of these vessels, running alongside dural venous sinuses, recapitulates the meningeal lymphatic system of rodents. In primates, meningeal lymphatics display a typical panel of lymphatic endothelial markers by immunohistochemistry. This discovery holds promise for better understanding the normal physiology of lymphatic drainage from the central nervous system and potential aberrations in neurological diseases.https://doi.org/10.7554/eLife.29738.001
How does the brain rid itself of waste products? Other organs in the body achieve this via a system called the lymphatic system. A network of lymphatic vessels extends throughout the body in a pattern similar to that of blood vessels. Waste products from cells, plus bacteria, viruses and excess fluids drain out of the body’s tissues into lymphatic vessels, which transfer them to the bloodstream. Blood vessels then carry the waste products to the kidneys, which filter them out for excretion. Lymphatic vessels are also a highway for circulation of white blood cells, which fight infections, and are therefore an important part of the immune system.
Unlike other organs, the brain does not contain lymphatic vessels. So how does it remove waste? Some of the brain’s waste products enter the fluid that bathes and protects the brain – the cerebrospinal fluid – before being disposed of via the bloodstream. However, recent studies in rodents have also shown the presence of lymphatic vessels inside the outer membrane surrounding the brain, the dura mater.
Absinta, Ha et al. now show that the dura mater of people and marmoset monkeys contains lymphatic vessels too. Spotting lymphatic vessels is challenging because they resemble blood vessels, which are much more numerous. In addition, Absinta, Ha et al. found a way to visualize the lymphatic vessels in the dura mater using brain magnetic resonance imaging, and could confirm that lymphatic vessels are present in autopsy tissue using special staining methods.
For magnetic resonance imaging, monkeys and human volunteers received an injection of a dye-like substance called gadolinium, which travels via the bloodstream to the brain. In the dura mater, gadolinium leaks out of blood vessels and collects inside lymphatic vessels, which show up as bright white areas on brain scans. To confirm that the white areas were lymphatic vessels, the experiment was repeated using a different dye that does not leak out of blood vessels. As expected, the signals observed in the previous brain scans did not appear.
By visualizing the lymphatic system, this technique makes it possible to study how the brain removes waste products and circulates white blood cells, and to examine whether this process is impaired in aging or disease.https://doi.org/10.7554/eLife.29738.002
Recent reports (Aspelund et al., 2015; Louveau et al., 2015b) described the existence of a network of true lymphatic vessels within the mammalian dura mater that runs alongside blood vessels, notably the superior sagittal and transverse sinuses. The dural lymphatic vessels display typical immunohistochemical markers that identify lymphatic vessels elsewhere in the body. They provide an alternate conduit for drainage of immune cells and cerebrospinal fluid (CSF) from the brain, beyond previously described pathways of flow: via arachnoid granulations into the dural venous sinuses, and via the cribriform plate into the ethmoid region (Weller et al., 2009). Although early reports, based on injections of India ink into the cisterna magna of the rat, suggested that the dural pathway accounts for only a minority of the drainage (Kida et al., 1993), the more recent studies (Aspelund et al., 2015; Louveau et al., 2015b), which are based on injections of fluorescent tracers and in vivo microscopy, indicate that the dural system may be substantially more important for drainage of macromolecules and immune cells than previously realized.
Whether a similar network of dural lymphatics is present in primates remains unknown. Moreover, noninvasive visualization of the dural lymphatics – a necessary first step to understanding their normal physiology and potential aberrations in neurological diseases – has not been reported. We therefore verified pathologically the existence of a dural lymphatic network in human and nonhuman primates (common marmoset monkeys) and evaluated two magnetic resonance imaging (MRI) techniques that might enable its visualization in vivo. First, the T2-weighted fluid-attenuation inversion recovery (T2-FLAIR) pulse sequence, which is the clinical standard for detecting lesions within the brain parenchyma, is highly sensitive to the presence of gadolinium-based contrast agents in the CSF (Mamourian et al., 2000; Mathews et al., 1999; Absinta et al., 2015). Second, ‘black-blood’ imaging sequences, which are typically used for measurement of vascular wall thickness or detection of atherosclerotic plaque, are tuned to darken the contents of blood vessels (even when they contain a gadolinium-based contrast agent), but in the process the images may highlight vessels with other contents and flow properties (Mandell et al., 2017). For comparison, we also acquired a postcontrast T1-weighted Magnetization Prepared Rapid Acquisition of Gradient Echoes (MPRAGE) MRI sequence, which is widely implemented for structural brain imaging and depicts avid enhancement of dura mater and blood vessels, but which would not be expected to discriminate lymphatic vessels.
Cerebral blood vessels have a highly regulated blood-brain barrier, protecting the neuropil from many contents of the circulating blood. Under physiological conditions, the blood-brain barrier prevents gadolinium-based chelates in standard clinical use from passing into the Virchow-Robin perivascular spaces and parenchyma, so that these structures do not enhance on MRI. On the other hand, dural blood vessels lack a blood-meningeal barrier, enabling leakage of circulating fluids and small substances, including gadolinium-based compounds. This explains the thin, though often incomplete, dural enhancement that is seen on conventional T1-weighted MRI scans under physiological conditions (Figure 1), as well as its abnormal diffuse or localized thickening in a variety of pathological conditions (Smirniotopoulos et al., 2007; Antony et al., 2015). Using high-resolution (in-plane resolution 270 × 270 μm or finer) T2-FLAIR and T1-weighted black-blood MRI images, obtained after the intravenous injection of standard FDA-approved contrast material (gadobutrol), we were able to visualize the collection of interstitial gadolinium within dural lymphatic vessels (maximum apparent diameter ~1 mm) in 5/5 human healthy volunteers and 3/3 common marmoset monkeys (Figure 1). Our results suggest that in the dura, similar to many other organs throughout the body, small intravascular molecules extravasate into the interstitium and then, under a hydrostatic pressure gradient, collect into lymphatic capillaries through a loose lymphatic endothelium (Sharma et al., 2008).
To further test this hypothesis, meningeal lymphatics were also assessed using a second gadolinium-based contrast agent, gadofosveset, a blood-pool contrast agent suitable for angiography (Lauffer et al., 1998). Gadofosveset binds reversibly to serum albumin, increasing its molecular weight from 0.9 to 67 kDa. Under physiological conditions, albumin has a low transcapillary exchange rate into the interstitial compartment, estimated to be on the order of 5% per hour, which explains the propensity of gadofosveset to remain within blood vessels (Richardson et al., 2015). In both species, gadofosveset did not reveal dural lymphatics, especially on T1-black blood images (Figure 2 and Figure 2—figure supplement 1). As expected, on T1-weighted MPRAGE images, gadofosveset provided superior intravascular enhancement, in both meningeal and parenchymal blood vessels, compared to gadobutrol (Figure 2).
On 3D-rendering of subtraction MRI images (Videos 1–2, Figure 1—figure supplement 1), dural lymphatics are seen running parallel to the dural venous sinuses, especially the superior sagittal and straight sinuses, and alongside branches of the middle meningeal artery. The topography of the meningeal lymphatics fits with the previously described network in rodents as well as our neuropathological data (Figures 3 and 4). It is worth noting that the lymphatics visualized by MRI are large slow-flow lymphatic ducts, whereas blind-ending and small lymphatic capillaries, clearly seen by histopathology (Figure 3 and Figure 3—figure supplement 1), are unlikely to be revealed by MRI. The induction of experimental autoimmune encephalomyelitis (EAE) did not affect detection of dural lymphatic vessel in either of the two animals that we tested (not shown).
To support our in vivo imaging results, we further investigated the existence and topography of lymphatics in coronal and longitudinal sections of human and marmoset dura mater. To accomplish this, we tested a variety of putative lymphatic endothelial markers and found that selective double immunostaining for D2-40 podoplanin/CD31 and for PROX1/CD31 was the most effective strategy in discriminating lymphatic vs. venous blood vessels in dura samples – a challenging task since lymphatics sprout from transdifferentiation of venous endothelium (Ny et al., 2005; Yaniv et al., 2006; Srinivasan et al., 2007; Aspelund et al., 2014; Lowe et al., 2015) and persistently share some endothelial markers. A branched network of lymphatics was clearly seen within the dura mater. On D2-40 podoplanin/CD31 double staining, we identified a total of 93 human dural lymphatics; most were collapsed, explaining the large range of maximum transverse diameters (range = 7–842 μm, mean = 125 μm, standard deviation = 161 μm). The density of dural lymphatics was higher around the venous sinuses than in more lateral areas of the dura, and higher within the meningeal layer than the periosteal layer of the dura. As expected, red blood cells were not seen within lymphatics. In marmosets, direct comparison between in vivo MRI and histopathology was performed (Absinta et al., 2014; Guy et al., 2016; Luciano et al., 2016). As shown in Figure 4 and Figure 4—figure supplement 1, the three dural vessels detected on coronal postcontrast T2-FLAIR and on subtraction images colocalize with three clusters of dural cells expressing the full panel of lymphatic endothelial markers (LYVE-1, D2-40 podoplanin, PROX1, COUP-TFII) and CCL21, a chemokine implicated in lymphatic transmigration.
In the ongoing debate about the precise localization of lymphatics within the meninges (either completely within the dura or ‘shared’ between the dura and the arachnoid) (Kipnis, 2016; Raper et al., 2016), our pathological data clearly show that at least some lymphatics are contained entirely within the dura (Figures 3 and 4). On limited evaluation, we were unable to visualize lymphatics within the leptomeninges, but additional dedicated studies, ideally with nonconventional tissue-preparation methods, are warranted to fully explore this possibility. A comprehensive map of the meningeal lymphatic network would have implications for unraveling the ways in which the meningeal lymphatics participate in waste clearance and in immune cell trafficking within the central nervous system (Louveau et al., 2015a; Kipnis, 2016; Raper et al., 2016).
In inflammatory pathological conditions, cellular migration toward the dural lymphatics might be profoundly enhanced by specific signaling and lymphatic plasticity (Alitalo et al., 2005; Kim et al., 2012; Stacker et al., 2014). Noteworthy in this context are clusters of extravascular CD3+ lymphocytes and CD68+ phagocytic meningeal macrophages that we observed in the dura of several multiple sclerosis autopsies (not shown), confirming intense immune cell trafficking and communication. Indeed, without proper normative comparison, we cannot rule out that the extensive presence of small lymphatics observed in multiple sclerosis dura samples is the result of inflammation-mediated lymphoangiogenesis. On the other hand, lymphatic dysfunction might impair waste clearance in neurodegenerative diseases and aging, in line with the recently captured deposition of β-amyloid in human dura in elderly people (Kovacs et al., 2016).
Differently from experiments implementing injections of tracers within brain structures, here we aimed primarily to image dural lymphatic vessels in human and nonhuman primates, but we could not prove whether dural lymphatic vessels drain immune cells, CSF, or other substances from the brain to deep cervical lymph nodes, nor could we assess any link with the glymphatic system (Iliff et al., 2012; Xie et al., 2013; Iliff et al., 2013). Such an analysis would probably require injection of specific MRI-detectable tracers and acquisition of MRI time-series (not thoroughly explored in the current work), an important future research direction for the nonhuman primate work.
Overall, our data clearly and consistently demonstrate the existence of lymphatic vessels within the dura mater of human and nonhuman primates. Together with recent studies in rodents, our results show that the meningeal lymphatic system is evolutionarily conserved in mammals and confirm, after exactly two centuries, what the Italian anatomist Paolo Mascagni speculated were lymphatic vessels at the surface of human brain (Mascagni and Bellini, 1816). The ability to image the meningeal lymphatics noninvasively immediately suggests the possibility of studying potential abnormalities in human neurological disorders.
We carried out human studies under a protocol (NCT02504840) approved by NIH Institutional Review Board. Informed consent was obtained from all participants. Formalin-fixed human brains were attained at autopsy after obtaining consent from the next of kin. Animal studies were performed under a protocol approved by the Institutional Animal Care and Use Committee.
We studied five healthy volunteers (three women, age range 28–53 years) and obtained scans on a 3-tesla MRI unit (Skyra, Siemens Healthcare, Erlangen, Germany), using the body coil for radiofrequency transmission and a 32-element phased array coil for reception. Human MRI analysis is described in more detail in Bio-protocol (Ha et al., 2018).
Prior to injection of gadolinium-based contrast agent, the following high-resolution MRI sequences were collected:
Whole-brain T1-Magnetization Prepared Rapid Acquisition of Gradient Echoes (MPRAGE, sagittal 3D turbo-fast low angle shot [TFL] sequence, acquisition matrix 256 × 256, isotropic resolution 1 mm, 176 slices, repetition time [TR]/echo time [TE]/inversion time [TI]=3000/3/900 ms, flip angle 9, acquisition time 5 min 38 s);
Limited T2-weighted Fluid Attenuation Inversion Recovery (FLAIR, coronal 2D acquisition over the superior sagittal sinus, field-of-view 256 × 256, 22 slices, reconstructed in-plane resolution 0.25 mm x 0.25 mm, 42 contiguous 3 mm slices, TR/TE/TI = 6500/93/2100 ms, echo train length 17, bandwidth 80 Hz/pixel, acquisition time 5 min), optimized for detection of gadolinium-based contrast agent in the subarachnoid space (Absinta et al., 2015);
Black-blood scan (coronal 2D acquisition, Sampling Perfection with Application optimized Contrasts using different flip angle Evolution [SPACE] sequence, field-of-view 174 × 174, matrix 320 × 320, reconstructed in-plane resolution 0.27 × 0.27 mm, 64 contiguous 0.5 mm sections, TR/TE = 938/22 ms, echo train length 35, bandwidth 434 Hz/pixel, acquisition time 7 min 50 s). A series of 2 or three overlapping coronal acquisitions were acquired to cover most of the cerebral hemispheres;
Whole-brain T2-FLAIR scan (coronal 3D acquisition, SPACE sequence, field-of-view 235 × 235, matrix 512 × 512, reconstructed in-plane resolution 0.46 × 0.46 mm, 176 1 mm sections, TR/TE/TI = 4800/354/1800 ms, nonselective inversion pulse, echo-train length 298, bandwidth 780 Hz/pixel, acceleration factor 2, acquisition time 14 min);
Whole-brain T1-SPACE (axial 3D acquisition, acquisition matrix 256 × 256, isotropic resolution 0.9 mm, 112 sections, TR/TE = 600/20 ms, flip angle 120, echo-train length 28, acquisition time 10 min).
We repeated these acquisitions after injection of gadobutrol (Gadavist, Bayer HealthCare, Whippany, NJ) in all five participants. In 2 of the participants, we also repeated the entire protocol prior to and following intravenous injection of gadofosveset (Lauffer et al., 1998) (Ablavar, Lantheus Medical Imaging, N Billerica, MA). Injections followed the manufacturer’s suggested dosing (0.1 mmol/kg for gadobutrol, 0.03 mmol/kg for gadofosveset).
We studied three healthy adult common marmosets (Callithrix jacchus) (one female, two males, age range 4–11 years). After the baseline MRI, experimental autoimmune encephalomyelitis (EAE) was induced in 2 marmosets with 0.2 mg of fresh-frozen human white matter homogenate as previously described (Gaitán et al., 2014). Marmosets were placed in a sphinx position within the magnet, and scans were obtained on a 7-tesla MRI unit (Avance AVIII, Bruker BioSpin, Billerica, MA, USA). Data acquisition was performed in transmit-only/receive-only mode using a homemade linear birdcage coil (120 mm inner diameter) as a radiofrequency transmitter and a homemade, 8-channel radio-frequency surface-array receiver coil assembly placed over the head of the animal. Prior to injection of contrast agent, we collected:
Whole-brain T1-weighted MPRAGE scan (coronal 3D acquisition, Modified Driven Equilibrium Fourier Transform [MDEFT] sequence, in-plane voxel size 0.15 mm x 0.15 mm, 36 contiguous 1 mm sections, TR/TE/TI = 12.5/4/1200 ms, flip angle 12 degrees, 2 segments of 1800 ms, acquisition time 7 min);
Whole-brain T2-FLAIR scan (coronal 2D acquisition, Rapid Acquisition with Relaxation Enhancement [RARE] sequence, voxel size 0.15 mm x 0.15 mm, 36 contiguous 1 mm sections, TR/TE/TI = 10,000/36/2500 ms, flip angle 90–180 degrees, 2 averages, acquisition time 13 min).
We performed the same scans following intravenous injection of gadobutrol and gadofosveset, in two different MRI sessions. Injections used single (0.1 mmol/kg for gadobutrol, 0.03 mmol/kg for gadofosveset) or triple the recommended human dosing (0.3 mmol/kg for gadobutrol, 0.09 mmol/kg for gadofosveset).
Scanner-generated DICOM images were converted into NIFTI files for postprocessing. Using MIPAV software (https://mipav.cit.nih.gov), precontrast scans were rigidly registered to postcontrast scans, and subtraction images were created for anatomical identification of dural lymphatic vessels. 3D skull-stripped subtraction images were then imported into Osirix software for maximum intensity projection (MIP) 3D rendering. The same postprocessing was performed for scans acquired after gadofosveset injection, and a direct comparison between the two gadolinium-based contrast agents was made.
Neuropathological evaluation focused on human and marmoset brain dura mater samples. Multiple human dura samples were obtained from 2 formalin-fixed brains (60- and 77-year-olds with longstanding progressive multiple sclerosis) and from a 33-year-old with refractory epilepsy undergoing anterior temporal lobectomy (Figure 3—source data 1). Detailed histopathological methods are described in Bio-protocol (Ha et al., 2018). Primate samples were obtained from 3 adult common marmosets (2 with EAE) [Figure 4—source data 1]. After general anesthesia and transcardial perfusion of 4% paraformaldehyde, marmoset brains were extracted and stored in 10% formalin. For primate brains, MRI-matched histological sections were achieved via 7T MRI of the fixed brain and subsequent gross sectioning with an individualized, MRI-designed, 3D-printed cutting box, as previously described (Absinta et al., 2014; Guy et al., 2016; Luciano et al., 2016).
Coronal and longitudinal 3 to 7 μm paraffin sections of the dura mater were stained with hematoxylin and eosin (H and E). Immunohistochemical and immunofluorescent analysis for classical lymphatic endothelial cell markers [lymphatic vessel endothelial hyaluronan receptor 1 (LYVE-1), podoplanin (D2-40), prospero homeobox protein 1 (PROX1), COUP transcription factor 2 (COUP-TFII) and CCL21], were performed on representative slides. Selective double staining for lymphatic vs. vascular endothelial markers (D2-40/CD31 and PROX1/CD31) were also executed on representative sections and implemented for histological quantification; the maximum diameter was recorded for each lymphatic vessel, and descriptive statistics were computed. We also stained for lymphocytes (CD3) and monocytes/macrophages (CD68). Five-μm paraffin sections of the human skin were used as positive controls for lymphatic endothelium cell marker assessment (Figure 3—figure supplement 2). For comparison, 10 paraffin human brain tissue blocks, selected for the presence of nearly intact leptomeningeal architecture, were implemented to assess the presence of lymphatics within the subarachnoid space and cerebral parenchyma.
Source, antibody type, and dilution are indicated sequentially as follows: LIVE-1 (Abcam, UK, rabbit polyclonal, 1:200); podoplanin D2-40 (AbD Serotec, Hercules, CA, mouse monoclonal, 1:50); PROX-1 (AngioBio, San Diego, CA, rabbit polyclonal, 1:50); COUP TFII (R&D Systems, Minneapolis, MN, mouse monoclonal, 1:200); CCL21 (Abcam, rabbit polyclonal, 1:200); CD31 (Abcam, rabbit polyclonal, 1:50); CD68 KP1 (Thermofisher, Waltham, MA, mouse monoclonal, 1:100); CD3 (Dako, Santa Clara, CA, rabbit polyclonal, 1:200); goat anti-mouse Alexafluor 488 IgG (Invitrogen, Carlsbad, CA, 1:250) and goat anti-rabbit Alexafluor 594 IgG antibodies (Invitrogen, 1:250).
Immunostaining was performed on 3 to 7 μm-thick consecutive paraffin sections with antibodies to LYVE1, D2-40 podoplanin, PROX1, COUP-TFII, CCL21, CD31, CD68, CD3 and visualized with 3,3'-diaminobenzidine (DAB). Briefly, after deparaffinization, sections were rinsed in triphosphate-buffered saline (TBS) for 10 min each, processed for antigen retrieval, and treated in 3% hydrogen peroxide for 10 min. Sections were blocked with a protein-blocking solution for 20 min and incubated with the primary antibodies overnight at 4°C or 1 hr at room temperature, then rinsed and incubated with secondary antibodies for 30 min. The immunoreaction was visualized with DAB. After washing, sections were counterstained with 10% hematoxylin. Stained sections were visualized and digitized (Axio Observer Z.1, Carl Zeiss Microscopy, NY, USA). Selective double staining for lymphatic and vascular endothelium markers (D2-40 podoplanin/CD31, PROX1/CD31), as well double staining for D2-40 podoplanin/PROX1,were also executed on representative sections. Double staining was performed using, in sequence, DAB horseradish peroxidase (HRP) and Vector Blue alkaline phosphatase (AP) methods. As negative control, sections of human skin, human dura mater, and marmoset brain were stained without the primary antibodies; no background staining and/or nonspecific binding was noted (Figure 3—figure supplement 2).
After deparaffinization, sections were treated using the same method as described above. Sections were incubated with a cocktail of antibodies to D2-40 podoplanin and CD31 for 2 hr at room temperature, then rinsed and incubated with secondary antibodies with Alexafluor 488 goat anti-mouse (Invitrogen, dilution 1:250) and Alexafluor 594 goat anti rabbit IgG antibodies (Invitrogen, dilution 1:250), respectively, for 30 min at room temperature. After washing, sections were mounted with DAPI Immuno Mount. Stained sections were visualized and digitized (Axio Observer Z.1, Carl Zeiss Microscopy, NY, USA).
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Heidi Johansen-BergReviewing Editor; University of Oxford, United Kingdom
In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.
Thank you for submitting your article "The primate meninges harbor lymphatic vessels that can be visualized noninvasively by MRI" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and David Van Essen as the Senior Editor.
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This report is a follow-up on the recent publications on the meningeal lymphatic vessels in rodents. They study image human and nonhuman primates using T2-FLAIR and T1-weighted imaging in combination with systemic administration of contrast agents. The analysis shows that the meningeal lymphatic vessels in human and nonhuman primate meninges are very similar to those described in rodents.
1) In this study, and in the previous work on the glymphatic system, there is little or no mention of the well-known Virchow-Robin perivascular spaces, easily observable using high resolution MRI (e.g. Thomas et al., MRM 2004). Because these features of brain anatomy are well known, having been studied for over a century, and they are also observable with MRI, it is quite surprising that they were not mentioned in the present manuscript. Were they not observed at all? If indeed they comprise a part of the lymphatic circulation, one would expect them to appear with significant contrast in the gadobutrol labelled images. The authors could provide additional visualisation.
2) Lymphatic flow. One of the criticisms of the 'glymphatic system' concept concerns the shortage of evidence that the lymph has a directional flow within it. A study of the kind described in this manuscript could help to rectify this lack – if a time series of images were obtained, displaying successive enhancement of lymphatic channels in the order predicted by the theory. The authors might want to comment on this possibility.
3) The MRI imaging is based on the use of two gadolinium-based tracers, one (gadofosveset) which is confined to the blood by binding to serum albumin, and one (gadobutrol) which can exit through blood vessel walls and pass into the lymph vessels. The latter showed accumulation in (putative) lymph vessels in the dura, while the former did not. This is encouraging but there are the following problems:
a) The putative lymphatics are barely visible, especially in the subtraction images in Figure 2. The authors should increase the intensity and the contrast of the images (taking care to specify what manipulations have been done in the figure legend) – for example, pasting Figure 2's subtraction images into Powerpoint and increasing both the intensity and contrast by 40% leads to much more easily visible lymphatics. A similar process should be applied to Figure 2—figure supplement 1 for the subtraction images, if the authors wish to convince the reader that there are no lymphatics visible with gadofosveset.
b) The videos show the extent of the lymphatics far more impressively than do the coronal sections in Figures 1 and 2. Panels should be added to Figures 1 and 2 showing sagittal sections or simply a picture of the 3D rendering in the videos. The contrast should also be increased in the videos.
4) Ultimately the MRI data need validating by immunocytochemistry with lymphatic specific markers, and this is attempted in Figures 3 and 4. Again there are problems:
a) Figure 3 is very poorly presented for readers not familiar with the detailed anatomy. I cannot understand what Figure 3B shows; add some explanatory labels. For panels D-E we need some clue as to exactly where in the brain the tissue is located – show a picture like that in Figure 4A, with rectangles to approximately locate the images presented in D-E. No scale bar size is stated for panels D and E and K.
b) In humans, Prox 1 or podoplanin (brown) are used as a lymphatic marker and CD31 (blue) as an endothelial marker. Unfortunately the images in panels 3D-E are so faint that colors are not visible (in contrast the immunofluorescence in panels J-K is much clearer). In the magnified panels colors are visible but the background labeling is high. Any dark brown label is attributed to the presence of lymphatics (e.g. there are 5 structures annotated LV in panel F) yet it is unclear how these vessels relate to those seen in the MRI pictures, where there are 3 lymph vessels in Figure 1. Were the IHC and the MRI done on the same brains? If so, is it not possible to state whether the number of lymph vessels found by MRI is the same as is found by IHC? The fact that the IHC finds more vessels (below the resolution of MRI?) at least needs to be discussed. Also, the fact that podoplanin is in cells other than lymph vessels should be mentioned (e.g. kidney podocytes and possibly capillary pericytes, which could confound the interpretation: https://www.ncbi.nlm.nih.gov/pubmed/25908104).
c) For the marmoset data, the text states that the number of lymphatics seen with MRI was the same as with IHC, yet the number of lymphatics seen in Figure 4A could be 2 or 3 (enhance the intensity and contrast, and label them with red arrows), while in Figure 4C It is unclear exactly which brownish structures are being defined as lymphatics – they need explaining better, e.g. by drawing a dashed red line round them.
d) For both human and marmoset tissue, it is essential to show the (hopefully absent) labeling when the primary antibody for PROX1 or D2-40 is omitted from the IHC. Ideally this should be in brain, but skin would be an acceptable substitute if brain is unfeasible.
e) Figure 3—figure supplement 2 and its associated text is fairly peripheral to the main point of this paper – it might be better in a separate paper combined with more rigorous data showing lymphatic trafficking of immune cells.https://doi.org/10.7554/eLife.29738.018
- Martina Absinta
- Seung-Kwon Ha
- Govind Nair
- Pascal Sati
- Nicholas J Luciano
- Kareem A Zaghloul
- Daniel S Reich
- Martina Absinta
- Martina Absinta
- Maryknoll Palisoc
- Stefania Pittaluga
- Antoine Louveau
- Jonathan Kipnis
- Antoine Louveau
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
The Intramural Research Program of NINDS supported this study. We thank the study participants, Irene Cortese and the NINDS Neuroimmunology Clinic for patient recruitment and care, the NIH Functional Magnetic Resonance Imaging Facility for MRI support, Afonso C Silva and his laboratory for access to the 7T animal scanner and customized hardware, Steven Jacobson and Emily Leibovitch for collaboration on our animal studies, and Roger Depaz for assistance in animal preparation.
Human subjects: We carried out human studies under a protocol (NCT02504840) approved by the NIH Institutional Review Board. Informed consent was obtained from all participants.
Animal experimentation: Animal studies were performed under a protocol approved by the Institutional Animal Care and Use Committee of the NIH (NIH Animal Studies Protocol 1308-15).
- Heidi Johansen-Berg, Reviewing Editor, University of Oxford, United Kingdom
This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.