Cellular characterization of the mouse collecting lymphatic vessels reveals that lymphatic muscle cells are the innate pacemaker cells

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Lymphatic System: Closing in on pacemaker cells

Pin-Ji Lei, Timothy P Padera

Insight Jul 22, 2025

eLife Assessment

This manuscript aims to identify the pacemaker cells in the lymphatic collecting vessels - the cells that initiate the autonomous action potentials and contractions needed to drive lymphatic pumping. Through the exemplary use of existing approaches (genetic deletions and cytosolic calcium detection in multiple cell types), the authors convincingly determine that lymphatic muscle cells are the origin of the action potential that triggers lymphatic contraction. The inclusion of scRNAseq and membrane potential data enhances a tremendous study. This fundamental discovery establishes a new standard for the field of lymphatic physiology.

https://doi.org/10.7554/eLife.90679.4.sa0

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Abstract
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Abstract

Collecting lymphatic vessels (cLVs) exhibit spontaneous contractions with a pressure-dependent frequency, but the identity of the lymphatic pacemaker cell is still debated. Here, we combined immunofluorescence and scRNAseq analyses with electrophysiological methods to examine the cellular constituents of the mouse cLV wall and assess whether any cell type exhibited morphological and functional processes characteristic of pacemaker cells. We employed inducible Cre mouse models to target-specific cell populations including CkitCreER^T2 to target interstitial cells of Cajal-like cells, PdgfrβCreER^T2 to target pericyte-like cells; PdgfrαCreER^TM to target CD34⁺ adventitial cells; and Myh11CreER^T2 to target lymphatic muscle cells (LMCs) directly. These inducible Cre lines were crossed to the fluorescent reporter Rosa26^mTmG, the genetically encoded Ca²⁺ sensor GCaMP6f, and the light-activated cation channel rhodopsin2 (ChR2). Only LMCs consistently, but heterogeneously, displayed spontaneous Ca²⁺ events during the diastolic period of the contraction cycle, and whose frequency was modulated in a pressure-dependent manner. Further, optogenetic depolarization with ChR2 induced propagated contractions only in LMCs. Membrane potential recordings in LMCs demonstrated that the rate of diastolic depolarization significantly correlated with contraction frequency. These findings support the conclusion that LMCs, or a subset of LMCs, are responsible for mouse cLV pacemaking.

Introduction

The spontaneous contractions of collecting lymphatic vessels (cLVs) are an integral component to fluid and macromolecule homeostasis as they provide the force to transport fluid from the interstitial spaces back to the blood circulation (Scallan et al., 2016). In humans, spontaneous contractile activity is estimated to account for 2/3 of lymph transport (Engeset et al., 1977), and this function is significantly compromised in patients suffering from lymphedema, whose cLVs typically display weak and irregular or entirely absent contractile activity (Olszewski, 2002). Ex vivo studies, in which the intraluminal pressure can be precisely controlled, have refined our understanding of the pressure-dependent regulation of contraction frequency (Benoit et al., 1989; Gashev et al., 2004), with some mouse cLVs displaying a 10-fold increase in contraction frequency over a 10 cmH₂O pressure gradient (Scallan and Davis, 2013; Zawieja et al., 2018a). The observation that cLVs, often cannulated at various lengths for ex vivo preparations, retain a consistently tunable contraction frequency points to the presence of (a) pacemaker cell(s) innate to the structure of the cLV wall and with a seemingly ubiquitous presence along the length of the vessel (Zawieja et al., 1993; Castorena-Gonzalez et al., 2018b). Furthermore, isolated cLVs typically display single pacemaker initiation sites unless damaged or electrically uncoupled by pharmacological inhibition of gap junctions or genetic deletion of Gjc1 (also known as Connexin 45, Cx45) (Behringer et al., 2017; Castorena-Gonzalez et al., 2018b; Castorena-Gonzalez et al., 2020). In sum, this suggests the pacemaker cell(s) is(are) likely both ubiquitous and continuous, to allow for electrical conduction via gap junctions, along the length of the cLV and prevent colliding contractile waves which would impair lymph transport.

Investigations into the cLV pacemaker identity have focused largely on cells termed interstitial cells of Cajal-like cells (ICLCs; or telocytes) (McCloskey et al., 2002; Briggs Boedtkjer et al., 2013), as they display some morphological and cell marker expression profiles similar to the interstitial cells of Cajal (ICC), which are bona fide pacemakers in the gastrointestinal (GI) tract. ICC are classically identified by either methylene blue staining and expression of CKIT, and coordinate GI smooth muscle contraction (Maeda et al., 1992; Ward et al., 1994; Ordög et al., 1999). ICC also expresses the canonical Ca²⁺-activated Cl⁻ channel Anoctamin 1 (Ano1) (Gomez-Pinilla et al., 2009), which is required for pacemaker activity (Hwang et al., 2009; Zhu et al., 2009; Singh et al., 2014). Previous reports in sheep mesenteric lymphatic vessels identified a population of CKIT⁺ and VIMENTIN⁺ ICLC in the vessel wall between the endothelial and lymphatic muscle cell (LMC) layer and running along the axis of the vessel (McCloskey et al., 2002). Investigations in the human thoracic duct also identified a significant population of ICLCs in close proximity to the LMCs evident by methylene blue staining, immunostaining for CD34, VIMENTIN, and CKIT, as well as the gold standard of electron microscopy (Briggs Boedtkjer et al., 2013). However, neither study could determine if these cells had functional electrical communication with the LMCs or demonstrate either a membrane electrical clock or internal Ca²⁺ clock to drive the rhythmic lymphatic vessel contractions observed ex vivo. LMCs share a functional similarity to ICC in that they also display the ANO1-mediated Ca²⁺-activated Cl⁻ current (van Helden, 1993; Toland et al., 2000; Mohanakumar et al., 2018; Zawieja et al., 2019), that regulates pacemaking. Spontaneous transient depolarizations, presumably ANO1 dependent, were recorded in mesenteric cLVs from guinea pigs (van Helden, 1993; von der Weid et al., 2008), providing a mechanism for membrane potential instability to drive AP initiation. Furthermore, computational models have proposed LMC sarcoplasmic reticulum (SR) Ca²⁺ release as the oscillator mechanism driving pacemaking (Imtiaz et al., 2007). SR Ca²⁺ release has also been implicated in pericyte regulation of arterioles (Hashitani et al., 2015; van Helden and Imtiaz, 2019), in microvascular vasomotion (Boedtkjer et al., 2008; Aalkjær et al., 2011; van Helden and Imtiaz, 2019), and in the contraction waves of atypical muscle cells of the lower urinary tract (Grainger et al., 2022).

Presently, no investigations have clearly identified the cellular identities of possible pacemaker cells within the cLVs of the mouse. Mouse cLVs exhibit contractile parameters and conduction speed equivalent to those of human vessels (Castorena-Gonzalez et al., 2018b) and their simplified architecture, compared to larger mammals, in combination with the genetic tools developed for the mouse model, allowed us to test for a fundamental pacemaker cell in the cLV. In this study, we utilized multiple genetic mouse models, confocal imaging of fluorescent reporters, cell-specific expression of GCaMP6f for Ca²⁺ imaging, and optogenetic light-activated depolarization to both visualize and test the functional aspects of putative pacemaker cells, along with membrane potential recordings in LMCs in pressure-challenged cLVs. We also performed immunostaining and single-cell RNA sequencing (scRNAseq) of isolated cLVs to provide greater detail to the heterogenous cellular populations found within the mouse cLVs. Despite identifying a significant population of CD34⁺PDGFRα⁺ adventitial cells along the length of mouse cLVs, the results of our functional studies support a myogenic (LMC) origin of pacemaking in cLVs.

Results

Methylene blue staining reveals a minor population of cells in mouse cLVs

Methylene blue staining was used to identify an ICLC population in the human lymphatic thoracic duct (Briggs Boedtkjer et al., 2013). In our isolated and cleaned lymphatic inguinal axillary collecting vessels (IALVs), methylene blue stained a significant number of cells with variable density along the length of the IALV and heterogeneous cell morphologies (Figure 1A–C). A significant portion of the stained cells resembled lymphatic vessel-associated macrophages with an elongated shape, while other cells were smaller and circular (Figure 1D–F). Methylene blue also appeared to stain mast cells as there were large ovoid cells with intracellular granules on the adventitial surface of the vessel. In addition, methylene blue stained a minor population of cells that exhibited long and thin axon-like extensions which appeared to have a slight helical orientation, with a small central body and nucleus (Figure 1C). None of these cell populations were aligned with the longitudinal axis of the vessel that would permit efficient coupling or regulation across the circumferential layer of LMCs required for coordinated propagation along the length of the vessel.

Figure 1

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Methylene blue staining of isolated mouse IALVs.

Representative image of an isolated and cleaned IALV after methylene blue staining which revealed cells of various morphology (A). (B) is the zoomed-in image of the yellow dotted box in A which contained large ovoid cells with granular staining (B, yellow asterisks). Fine cellular extensions (red asterisks) stained by methylene blue in some cells were visualized with color channel separation and division (C). (**D, E**) Similar to B and C, but in a separate vessel which stained with a higher density of methylene blue stained cells, some of which had limited cellular processes. (F) Focal reconstruction from imaging a methylene blue stained IALV using an upright microscope and immersion objective. Methylene blue staining was performed in IALVs isolated from five mice (n = 5).

Immunofluorescence Imaging of IALVs Stained for ICLC, LEC, and LMC Markers

We next stained IALVs for the putative telocyte/ICLC markers CKIT, CD34, and the intermediate filament VIMENTIN, which have been previously utilized for ICLC identification in human and sheep lymphatic tissues (McCloskey et al., 2002; Briggs Boedtkjer et al., 2013). Additionally, an antibody to the intermediate filament DESMIN was used to label muscle cells (McCloskey et al., 2002). IALVs stained for cKIT (Figure 2B) showed robust signal in large ovoid cells with a non-segmented circular nucleus (Figure 2A), characteristic of mast cells that were in the outer part of the adventitia. Similarly, CKIT-stained populations of elongated cells as well as circular cells with variable densities were observed throughout the IALV wall, similar to methylene blue⁺ cell populations (Figure 2B, J). Staining for CD34 revealed a large population of cells that were seemingly contiguous along the length of the vessel. The CD34⁺ cells generally had multiple lobular processes and a ‘oak leaf’-like appearance, typically a characteristic of fibroblasts, though some contained short, thin dendrite-like extensions (Figure 2C, G, K). The CD34⁺ cells were negative for DESMIN (Figure 2H), which primarily stained the circumferential LMCs (Figure 2F). Note that the largely non-circumferential cell organization in this region is typical for a lymphatic endothelial valve site (Bridenbaugh et al., 2013a). Furthermore, CD34⁺ and cKIT⁺ cells were separate populations (Figure 2D, L). A VIMENTIN antibody labeled lymphatic endothelial cells (LECs) which exhibited a horizontal cobblestone morphology in parallel with the vessel axis (Figure 2E, I), while also co-labeling the majority of the CD34⁺ cells (Figure 2H) and CKIT⁺ cells (Figure 2L). Videos of the half vessel z-stacks are provided (Figure 2—video 1; Figure 2—video 2; Figure 2—video 3 for Figure 2D, H, L, respectively).

Figure 2 with 3 supplements see all

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Staining mouse IALVs for ICLC markers.

Representative immunofluorescent max projections of half vessel confocal image stacks imaged from mouse IALVs stained for ICLC markers. DAPI (A), cKIT (B), and CD34 (C) and their merged image (D). Representative max projections of the intermediate filament VIMENTIN (E), the intermediate filament DESMIN (F), CD34 (G), and their merged image (H). Representative max projection of VIMENTIN (I), CKIT (J), CD34 (K), and their merged image (L). Scale bar = 100 µm for all images.

Of the cells stained in Figure 2, the CD34⁺ population was intriguing due to its high density and distribution throughout the length of the IALV, which potentially would be conducive to effective regulation of LMC excitability. In addition to CD34 and VIMENTIN, PDGFRα staining is also commonly ascribed to both telocytes (Vannucchi et al., 2013; Xiao et al., 2013; Zhou et al., 2015) as well as fibroblasts (Kimura et al., 2021; Clayton et al., 2022). We performed immunofluorescence imaging for PGDFRα counterstained with CD34 and markers for LMCs, LECs, and pericytes. As noted in Figure 2, CD34⁺ cells (Figure 3A) did not co-label LMCs (Figure 3D), which were smooth muscle actin⁺ (ACTA2, Figure 3B) and CALPONIN⁺ (Figure 3C). However, nearly all CD34⁺ (Figure 3E) cells were also PDGFRα⁺ (Figure 3F, H). Occasionally, some overlap of PDGFRα and ACTA2 signal was noted (Figure 3G, H). LECs staining with PECAM1 (Figure 3I) revealed the expected rectangular elongated cobblestone morphology that was distinct from the PDGFRα⁺ cells (Figure 3J, L). Staining for CALPONIN also specifically labeled LMCs (Figure 3K) but not PDGFRα⁺ cells (Figure 3L). Lastly, we stained for PDGFRα, CD34, and the commonly used pericyte marker PDGFRβ (Figure 3M–P). As above, CD34 and PDGFRα were highly colocalized (Figure 3Q, R, T), and many of the CD34⁺ and PDGFRα⁺ cells were also PDGFRβ⁺ (Figure 3P). PDGFRβ also stained some circumferential LMCs (Figure 3Q). During the imaging of mouse IALVs for these markers, we also observed that the lymphatic secondary endothelial valves were populated by elongated cells that stretched the length of the valve leaflet and were positive for CD34, PDGFRα, and PDGFRβ, with varying intensities. These cells could be observed in most, if not all, the valves we assessed and found within both leaflets of the valve (Figure 3R, S). These cells had long, thin extensions that were branched, along with apparent dendrite-like extensions with a morphology that closely resembled those described for pericytes or telocytes (Popescu and Faussone‐Pellegrini, 2010). PDGFRα⁺ or CD34⁺ cells with this morphology were only observed in the valve leaflets, and thus seemed insufficient to regulate pacemaking as normal contractions are observed in isolated cLVs without secondary valves (van Helden, 1993; Gashev et al., 2002). Representative z-stacks demonstrating these valve-located ‘telocyte’ shaped cells (Figure 3R, S) are provided in Figure 3—videos 1 and 2.

Figure 3 with 4 supplements see all

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Immunofluorescence labeling of mouse IALVs with markers for ICLC, lymphatic muscle cell (LMC), lymphatic endothelial cell (LEC), and immune cell populations.

We stained isolated mouse IALVs with cellular markers used to differentiate various cell types observed in collecting lymphatic vessels (cLVs). Half vessel image stacks were taken with confocal microscopy and the resulting representative max projections are shown. (A) CD34 stained cells and LMC staining with ACTA2 (B) and CALPONIN (C) and the corresponding merged (D) image. There was significant overlap in (E) CD34 staining along with the fibroblast marker PDGFRα (F) compared to LMC staining with ACTA2 (G) and the merged (H) image. The endothelial marker PECAM1 (I) to delineate LECs with PDGFRα staining (J), and the LMC marker CALPONIN (K) with the merged image (L) revealed three separate populations of cells. PDGFRβ (O) stained many cells that were CD34 (M) and PDGFRα (N) positive, as seen in the merge imaged (P), in addition to PDGFRβ signal detected in the LMC layer (Q). Max projections of only the luminal frames of a z-stack at lymphatic valve locations revealed PDGFRβ, CD34, and PDGFRα labeling in bipolar shaped cells with long extensions that traveled throughout the valve leaflets (R, S). Control IALV (T) stained only with secondary antibody. Scale bar = 100 µm for all images.

We next determined the degree of colocalization between the CD34 and PDGFRα signal given the significant overlap in their staining profile. Colocalization analysis of PDGFRα (Figure 3—figure supplement 1A) and CD34 (Figure 3—figure supplement 1B) and their colocalization (Figure 3—figure supplement 1C) was determined with the FIJI BIOP-JACoP tool. The Pearson’s coefficient was 0.83 (Figure 3—figure supplement 1D) and Mander’s coefficient of overlap was 0.80 for the PDGFRα⁺ signal and 0.87 for the CD34 signal (Figure 3—figure supplement 1E). Colocalization between MYH11 and PDGFRα was significantly lower (Figure 3—figure supplement 1D–F) with a Pearson’s coefficient of 0.30 (Figure 3—figure supplement 1G), whereas the Mander’s coefficient for MYH11 overlap with PDGFRα was 0.077 and 0.043 for PDGFRα signal overlap with MYH11 (Figure 3—figure supplement 1H). The high degree of colocalization of CD34 and PDGFRα signal informed our use of the commercially available transgenic PdgfrαCreER^TM mouse model to target these cells. The vast majority of the PDGFRα⁺ cells were located in the adventitial layer (Figure 3—figure supplement 2A–D), which varied between 1 and 3 PDGFRα⁺ cells thick (Figure 3—figure supplement 2E). Under this layer, we observed only a single layer of largely circumferential LMCs stained by MYH11 (Figure 3—figure supplement 2B) sitting atop a single layer of PECAM1⁺ LECs (Figure 3—figure supplement 2A). We also observed occasional PDGFRα⁺ cells or their extensions located in the sub-endothelial space (Figure 3—figure supplement 2E’, E”) positioned between the LECs and the LMCs.

Use of inducible Cre-mediated recombination of Rosa26^mTmG to delineate and characterize specific IALV cell types

After confirming the presence of VIMENTIN⁺, cKIT⁺, and CD34⁺ PDGFRα⁺ positive cells within the mouse IALV, we sought to further investigate these cell populations by using constitutive and inducible Cre recombinase expressing mouse lines. IALVs from the constitutively active PdgfrαCre-Rosa26^mTmG and Cspg4Cre-Rosa26^mTmG mice had GFP fluorescence in the majority of LMCs as well as in the fibroblast-shaped cells found within the IALV wall (Figure 4A, B). While informative of expression of the LMC progenitor cells, neither constitutive Cre would be useful in delineating cell types. In contrast to the constitutively active PdgfrαCre, the tamoxifen-inducible PdgfrαCreER^TM line drove significant recombination in only the fibroblast-shaped cells previously stained for CD34 and PDGFRα but not in LMCs or LECs (Figure 4C). PdgfrβCreER^T2, commonly used to label pericytes, drove recombination in both a minor population of the LMCs and the fibroblast-shaped cells. CkitCreER^T2, which capably drives recombination in the ICCs of the GI tract (Baker et al., 2016), drove recombination only in a small population of irregularly spaced, large ovoid cells on the surface of the IALV (Figure 4E), although recombination in one or two LECs could occasionally be detected (not shown). Finally, Myh11CreER^T2 drove recombination in nearly all LMCs, which were largely circumferentially oriented with dendrite-like, cell-cell contacts visible between them and without significant GFP fluorescence in either LECs or the fibroblast-shaped CD34⁺ PDGFRα⁺ cell population (Figure 4F). Additionally, some LMCs maintained the bipolar shape but had secondary extensions forming a ‘Y’ shape in which an adjacent LMC typically filled the inner void. A very minor population of recombined cells in the Myh11CreER^T2-Rosa26^mTmG IALVs was smaller and irregularly patterned with multiple fine axon-like projections or ruffled edges (Figure 4F).

Figure 4

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Inducible Cre-*Rosa26^mTmG* labeling and fidelity to target putative pacemaker cell populations.

Stitched montages of serial max projections of GFP and tdTomato signal from live IALVs isolated from PdgfrαCre-*Rosa26^mTmG* (A), Cspg4Cre-*Rosa26^mTmG* (B), PdgfrαCreER^TM-*Rosa26^mTmG* (C), PdgfrβCreER^T2-*Rosa26^mTmG* (D), CkitCreER^T2-*Rosa26^mTmG* (E), and Myh11CreER^T2-*Rosa26^mTmG* (F). IALVs were digested into single cells and GFP⁺ cells were purified via FACS from Prox1-eGFP (G), Myh11CreER^T2-*Rosa26^mTmG* (H), PdgfrαCreER^TM-*Rosa26^mTmG* (I), and PdgfrβCreER^T2-*Rosa26^mTmG* (J) mice. Representative gels demonstrating RT-PCR products corresponding to the respective genes used in the promoter of each specific transgene employed to drive either eGFP or Cre-mediated recombination of *Rosa26^mTmG* from each GFP⁺-sorted population (K–N) to assess fidelity. Images are representative of IALVs from at least three separate mice (n = 3). FACS and RT-PCR were repeated at least three times (n = 3 mice).

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To complement the morphological and cell density findings obtained with confocal microscopy, we digested IALVs from the inducible Cre-Rosa26^mTmG lines, and the Prox1-eGFP line as a control, into single-cell suspensions and sorted the respective GFP⁺ populations (Figure 4G–J) for RT-PCR profiling (Figure 4K). We first focused on determining the molecular fidelity of the sorted cells based on the gene promoters used to drive each inducible Cre model to discern cellular overlap. In agreement with the confocal images, sorted GFP⁺ cells from PdgfrβCreER^T2-Rosa26^mTmG IALVs expressed Pdgfrb but also Myh11 and Pdgfra. In contrast, GFP-sorted cells from PdgfrαCreER^TM IALVs expressed Pdgfra and Pdgfrb, but with no detectable expression of Myh11. GFP⁺ cells from sorted Myh11CreER^T2-Rosa26^mTmG IALVs had high expression for Myh11 as well as Pdgfrb, but did not express Pdgfra. IALVs from CkitCreER^T2-Rosa26^mTmG mice were not pursued for FACS due to the exceptionally sparse recombination observed along the IALV.

Characterization of the cellular components of the mouse IALVs by scRNAseq and FACS–RT-PCR

The results from the immunofluorescence staining, Rosa26^mTmG reporter imaging, and FACS–RT-PCR experiments suggested that both LMCs and AdvCs can express Pdgfrb. To provide further clarity and detail to the cellular populations within the mouse cLV wall and potential subsets within those broad cell types, we performed scRNAseq on isolated and cleaned inguinal axillary cLVs from male and female mice. The resulting uniform manifold approximation and projection (UMAP) (Figure 5A) revealed a host of cell types which had three main clusters corresponding to LECs, LMCs, and AdvCs (Figure 5A). We assessed the expression of genes that correspond to the markers from our earlier immunofluorescence staining as well as cell identification markers commonly used within the literature to identify each cell cluster (Figure 5B). Cell identity was confirmed by commonly used markers (Figure 5B) and the top differentially expressed genes (Figure 5—figure supplement 1A). Feature plots for the LEC markers Prox1 (Figure 5C) and Flt4 (Figure 5D), LMC markers Myh11 (Figure 5E) and Cnn1 (Figure 5F), and the AdvCs markers Pdgfra (Figure 5G) and Lumican (Figure 5H) were quite specific for labeling their respective cell clusters. Very few Kit (Figure 5I) expressing cells were observed in accordance with our imaging results. Pdgfrb was observed in both LMC and AdvC clusters (Figure 5J) while the remaining cell clusters were of immune origin as they expressed the gene Ptprc encoding the hematopoietic marker CD45 (Figure 5K). Notably, the previous genes suggested to identify LMCs in a previous scRNASeq study (Kenney et al., 2022), Dpt, Pi16, and Ackr3, were largely absent in LMCs and instead were expressed in a minor population of AdvCs (Figure 5—figure supplement 1B). We provide a further sub-clustering breakdown of the LECs (Figure 5—figure supplement 2), LMCs (Figure 5—figure supplement 3), AdvCs (Figure 5—figure supplement 4), and a detailed expression profile of the immune cell clusters (Figure 5—figure supplement 5). Further assessment of the LEC subcluster included a putative lymphatic endothelial ‘up valve’ cell population in subcluster 8 which expressed high levels of Prox1, Cldn11, Itga9, Gja4, and Neo1 and ‘down-valve’ population in cluster 6 which expressed Clu, Adm, Gja4, and Lypd6 (Figure 5—figure supplement 2C) which mapped well to previous RNAseq datasets (González-Loyola et al., 2021; Petkova et al., 2023; Yoon et al., 2024; Takeda et al., 2019). The top differentially expressed genes in the putative up-valve population in cluster 8 included Irx3, Neo1, Tub, Ano4, and Fxyd2, and we noted Cacna1e, Fgf14, and Irf1 in the down-valve cluster 6. Analysis of the LMC subclusters did not reveal any significant differences in the expression of known pacemaking-associated genes Ano1 or Itpr1 at our initial conditions of Log2FC of 0.5. However, we provide an overview of the typical ion channel families expressed in LMCs in Figure 5—figure supplement 3B–I. The AdvC cells could be further subclustered into multiple populations (Figure 5—figure supplement 4A and C) with little evidence of LMC gene contamination as these cells lacked Myh11, Kcnma1, and Tagln despite expression of Cacna1c, Ano1, and Gjc1. Over 75% of AdvCs expressed Pdgfra (Figure 5—figure supplement 4D) and 65% of the total AdvCs co-expressed both Pdgfra and Cd34. Our immunofluorescent colocalization of PDGFRα and CD34 staining was also supported as 72% of AdvCs expressing either Cd34 or Pdgfra co-expressed both genes (Figure 5—figure supplement 5D). The vast majority of AdvCs expressing Pdgfrb (Figure 5—figure supplement 4E) or Cspg4 (Figure 5—figure supplement 4F) also expressed Pdgfra. Expression of Ano1, Gjc1, and Cacna1c was also observed in some of the AdvCs, and most of those cells also co-expressed Pdgfra, supporting further use of the PdgfrαCreER^TM line (Figure 5—figure supplement 4G–I).

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scRNAseq analysis of mouse IALVs from *Rosa26^mTmG* mice.

IALVs were cleaned and isolated from 10 *Rosa26^mTmG* mice and digested into a single-cell suspension for scRNAseq analysis with the 10X platform. (A) Uniform manifold approximation and projection (UMAP) of the various cell populations that compromise the mouse IALV, though some mammary epithelia contamination was present (populations 18 and 19). (B) Heatmap of commonly used genes for cell identification for each of the cell clusters. Feature plots to assess cell cluster expression of the genes shown in Figure 4 inlcuding the lymphatic endothelial cell (LEC) markers *Prox1* (C) and *Flt4* (D), lymphatic muscle cell (LMC) markers *Myh11* (E) and *Calponin 1* (F, *Cnn1*), fibroblast markers *Pdgfra* (G) and *Lum* (H, *Lumican*), ICC marker *Kit* (I), the pericyte and smooth muscle precursor marker (*Pdgfrb*) (J), and the hematopoietic marker *Ptprc* (K).

While scRNAseq highlighted the depth of heterogeneity of the cellular composition of the mouse cLV, we wanted to validate the actual recombined cell populations from the inducible Cre-Rosa26^mTmG lines. We profiled each inducible Cre-driven recombination of Rosa26^mTmG via FACS-purified cells and RT-PCR for common markers for endothelial cells, muscle cells, and pericytes. Nos3 (eNOS) expression was observed only in the Prox1-eGFP sorted cells, and LECs also expressed Vim, Mcam, and had weak but detectable signal for Cd34 (Figure 6A). Myh11CreER^T2 sorted cells showed expression of smooth muscle actin (Acta2), the alpha subunit of the L-type voltage-gated Ca²⁺ channel Cacna1c (Cav1.2), Desmin (Des), Mcam, and Vimentin (Vim, Figure 6B). In addition to the genes expressed in Myh11CreER^T2 recombined cells, Cdh5, Cd34, and Cspg4 (also known as Ng2) were also detected in cells sorted from PdgfrβCreER^T2 IALVs (Figure 6C). As expected, the GFP⁺ cells sorted from PdgfrαCreER^TM IALVs expressed mRNA for Cd34, weak signal for Cspg4, and Vimentin, but not Desmin, Acta2, nor the pericyte marker Mcam (Figure 6D). Cacna1c was expressed in cells FACS purified from both PdgfrβCreER^T2 and Myh11CreER^T2 IALVs and sorted cells from PdgfrαCreER^TM IALVs without any evidence that Myh11-expressing muscle cells contaminated the latter. These findings confirmed the separate cell populations achieved with PdgfrαCreER- and Myh11CreER^T2-mediated recombination, at least as it pertains to the Rosa26^mTmG reporter. These findings were largely validated by our scRNASeq dataset. Cdh5 (Figure 6E) and Nos3 (Figure 6F) were almost exclusively expressed in the LEC clusters, while Acta2 (Figure 6G) was highly expressed in the LMC cluster. We also observed that Cacna1c was highly expressed in the LMCs (Figure 6H) and some AdvCs. Cd34 was widely expressed in AdvCs matching our immunofluorescence data. Cd34 expression was also seen in LECs (Figure 6I) although we did not observe a signal in LECs in our earlier immunofluorescence staining (Figure 3). Cspg4 was observed in a minor population of AdvCs (Figure 6J). The intermediate filament Vim (Figure 6K) was ubiquitously expressed across all clusters expressed, but Des was primarily expressed in LMCs and some subsets of AdvCs (Figure 6K, L). The endothelial and pericyte marker Mcam (also referred to as Cd146) was expressed in LECs and LMCs but was largely absent in AdvCs (Figure 6M). We followed up the identification of Cacna1c expression in the PdgfrαCreER^TM sorted cell population by assessing the expression of other genes involved in either electrical conduction (Gjc1) (Figure 6N) or pacemaking (Ano1) (Figure 6O) of IALVs. Expression of Ano1 and Gjc1 was observed in PdgfrαCreER^TM Rosa26^mTmG FACS-purified cells (Figure 6P).

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RT-PCR Profiling of FACS-purified cells from inducible Cre-*Rosa26^mTmG*.

Expanded RT-PCR profiling of genes to discriminate lymphatic endothelial cells (LECs), lymphatic muscle cells (LMCs), and other cell types in our GFP⁺ sorted cells from Prox1-eGFP (A), Myh11CreER^T2-*Rosa26^mTmG* (B), PdgfrβCreER^T2-*Rosa26^mTmG* (C), and PdgfrαCreER^TM-*Rosa26^mTmG* (D). Feature plots for the genes assessed in A-D in our IALV scRNAseq analysis confirmed those results (**E-M**). In addition to a population of AdvCs expressing *Cacna1c*, we also noted expression of *Gjc1* (N), which was also observed in LECs, and *Ano1* (O) in the AdvC clusters. We confirmed this expression using GFP⁺ cells sorted from PdgfrαCreER^TM-*Rosa26^mTmG* IALVs for RT-PCR (P) and ruled out hematopoietic or LEC contamination. All RT-PCRs were performed two to four times for each gene over each sorted cell population collected from different mice.

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Inducible deletion of either Cacna1c, Ano1, or Gjc1 with PdgfrαCreER^TM did not affect cLV pacemaking

The expression of the genes critically involved in cLV function—Cacna1c, Ano1, and Gjc1—in the PdgfrαCreER^TM-Rosa26^mTmG purified cells and scRNAseq data prompted us to generate PdgfrαCreER^TM-Ano1^fl/fl, PdgfrαCreER^TM-Gjc1^fl/fl, and PdgfrαCreER^TM-Cacna1c^fl/fl mice for contractile tests. We isolated popliteal cLVs and tested their pacemaker and contractile function in response to a physiological pressure range of 0.5–10 cmH₂O, under normal conditions. However, we did not detect any significant differences in pacemaking or contractile function as assessed by contraction frequency, ejection fraction, and vessel tone in popliteal cLVs studied from PdgfrαCreER^TM-Ano1^fl/fl mice (Figure 7A–C) or PdgfrαCreER^TM-Gjc1^fl/fl mice (Figure 7D–F). There was no difference in contraction frequency of cLVs from PdgfrαCreER^TM-Cacna1c^fl/fl mice compared to floxed control mice; however, we noted a mild but statistically significant increase in ejection fraction at the lowest pressure, 0.5 cmH₂O (Figure 7H). Additionally, vessels isolated from PdgfrαCreER^TM-Cacna1c^fl/fl mice also had a statistically significant increase in vessel tone (Figure 7I) noted at the two-way level although we did not resolve significance at any specific pressure with this sample. No differences in normalized contraction amplitude, fractional pump flow, or diastolic diameter were observed (Figure 7—figure supplement 1). In total, despite the presence of transcript for these critical genes in Pdgfra expresing cells, PdgfrαCreER^TM-mediated deletion of Gjc1, Cacna1c, or Ano1 failed to recapitulate previous reports of the significant contractile defects using the Myh11CreER^T2 line to delete the same genes (Castorena-Gonzalez et al., 2018b; Zawieja et al., 2019; To et al., 2020; Davis et al., 2022).

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Isobaric contractile assessment of popliteal collecting lymphatic vessel (cLV) from PdgfrαCreER^TM-driven deletion of *Ano1*, *GJC1*, and *Cacna1c*.

Summary of the contractile parameters recorded from popliteal cLVs in PdgfrαCreER^TM-*Ano1^fl/fl*, PdgfrαCreER^TM-*Gjc1^fl/fl* mice, PdgfrαCreER^TM-*Cacna1c^fl/fl* mice. Contraction frequency (A, D, G), ejection fraction (B, E, H), and vessel tone (C, F, I) were assessed. No statistically significant differences were observed in cLVs isolated from PdgfrαCreER^TM-*Ano1^fl/fl* and PdgfrαCreER^TM-*Gjc1^fl/fl* mice across these three parameters. Mean and SEM shown, n = 6 popliteal vessels from three mice PdgfrαCreER^TM-*Ano1^fl/fl* mice and n = 9 popliteal vessels from six mice *Ano1^fl/fl* mice. Mean and SEM shown, n = 5 popliteal vessels from three mice PdgfrαCreER^TM-*GJC1^fl/fl* mice and n = 11 popliteal vessels from eight mice *GJC1^fl/fl* mice. Mean and SEM shown, n = 6 popliteal vessels from three mice PdgfrαCreER^TM-*Cacna1c^fl/fl* mice and n = 8 popliteal vessels from eight mice *Cacna1c^fl/fl* mice. The contractile data from control *Cacna1c^fl/fl* vessels is a subset of previously published data that was separated by sex (Davis et al., 2022) while they are combined here. * denotes significance at p < 0.05 which 0.10 > p > 0.05 are reported as text. Normalized contraction amplitude, fractional pump flow, end diastolic diameter can be found in Figure 7—figure supplement 1.

PDGFRα⁺ adventitial fibroblasts express markers associated with multipotency

Despite the lack of cLV pacemaking deficits in the PdgfrαCreER^TM genetic knockout lines, we were curious to discern further insight into the role or function of the PDGFRα⁺ CD34⁺ cells since they comprise a significant portion of the lymphatic cLV wall. We performed RT-PCR on FACS-purified cells from Prox1-eGFP, Myh11CreER^T2-Rosa26^mTmG, and PdgfrαCreER^TM-Rosa26^mTmG IALVs for multipotency markers including Krüppel-like factor 4 (Klf4), lymphocyte antigen 6 family member A (Ly6a) (also referred to as stem cell antigen 1, Sca1), and Gli1, with Cd34 and Pdgfra used to assess purity. Recombined (GFP⁺) cells from Myh11CreER^T2-Rosa26^mTmG had weak expression of Klf4 and Gli1 but were negative for Ly6a (Figure 6—figure supplement 1A). PdgfrαCreER^TM recombined cells strongly expressed Klf4, Ly6a, and Gli1 (Figure 6—figure supplement 1A). LECs sorted from Prox1-eGFP IALVs were positive for Klf4, weak for Ly6a, and positive for Cd34 but negative for Gli1 and Pdgfra (Figure 6—figure supplement 1B). The unrecombined population (tdTomato⁺) cells in the Myh11CreER^T2- Rosa26^mTmG IALVs (Figure 6—figure supplement 1B) showed expression for all the markers as expected. PdgfrαCreER^TM recombined cells also expressed the mesenchymal stromal cell markers Itgb1, Eng, and Cd44 (Figure 6—figure supplement 1C, positive control in 9D). However, expression of these genes was not homogenous across all the AdvCs population based on our scRNAseq analysis (Figure 6—figure supplement 1E–J). We performed immunofluorescence staining for one of these multipotent markers, LY6A (Figure 6—figure supplement 1K) in the adventitial cells with PDGFRα (Figure 6—figure supplement 1L) and counter staining for LMCs with MYH11 (Figure 6—figure supplement 1M). The morphology and staining pattern of LY6A overlapped significantly with PDGFRα staining and not MYH11 staining (Figure 6—figure supplement 1N, Figure 6—video 1).

Optogenetic stimulation of inducible Cre-driven channel rhodopsin 2

We next used optogenetic methods to test whether the cell populations recombined by either CkitCreER^T2, PdgfrαCreER^TM, or Myh11CreER^T2 could elicit a coordinated contraction. The ChR2-tdTomato construct appeared more sensitive to recombination than Rosa26^mTmG, in some cases resulting in LMC expression of ChR2-tdTomato in PdgfrαCreER^TM and CkitCreER^T2 popliteal cLVs based on cell morphology. Care was taken to image each vessel for tdTomato (Figure 8A, C, E) prior to stimulation at its respective sites under brightfield conditions for diameter tracking (Figure 8B, D, F) to ensure fidelity of the cell types and morphologies observed in Figures 3 and 4. As with Rosa26^mTmG, CkitCreER^T2 drove the ChR2-tdTomato expression primarily in large ovoid cells found on the adventitia of the vessel. Cells were stimulated by positioning an optical laser fiber (tip diameter 2–3 μm) near a ChR2+ cell, with an illumination field of 10–50 μm. Localized photo-stimulation of these cells did not initiate coordinated contractions (Figure 8G–J, S). Similarly, photo-stimulation of ChR2-tdTomato expressing cells driven by PdgfrαCreER^TM failed to initiate a coordinated contraction (Figure 8K–N, T). In contrast, localized photo-stimulation of LMCs, using Myh11CreER^T2 to express Chr2-tdTomato, resulted in a propagated contraction in the popliteal vessel (Figure 8O–R, U). In total, only 3.25% of photo-stimulation events for CkitCreER^T2-ChR2-TdTomato and 3.03% of photo-stimulation events for PdgfrαCreER^TM-ChR2-tdTomato were associated with a contraction, while 88.5% of photo-stimulation events for Myh11CreER^T2-ChR2-tdTomato induced contractions (Figure 8V). The optogenetic triggering of contractions observed in PdgfrαCreER^TM-ChR2-tdTomato and CkitCreER^T2-ChR2-TdTomato vessels is likely due to the happenstance of spontaneous contractions occurring during the time and proximity of optogenetic stimulation (see Methods). As a control, we also used non-induced (no tamoxifen) Myh11CreER^T2-ChR2-tdTomato cLVs and contractions were associated with only 7% of photo-stimulation events, in line with the PdgfrαCreER^TM and CkitCreER^T2 results (Figure 8V). As mast cells are not ascribed to any tissue-specific pacemaking behavior, these similar low percentages observed between these three groups are suggestive of random coincidence. Brightfield videos of the photo-stimulation and representative traces for CkitCreER^T2-ChR2-TdTomato, PdgfrαCreER^TM-ChR2-tdTomato, and Myh11CreER^T2-ChR2-tdTomato are provided in Figure 8—video 1; Figure 8—video 2; Figure 8—video 3.

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ChR2-mediated depolarization only in lymphatic muscle cells (LMCs) triggers contraction.

Representative max projections of tdTomato-ChR2 signal in popliteal collecting lymphatic vessels (cLVs) isolated from CkitCreER^T2-ChR2-tdTomato (A), PdgfrαCreER^TM-ChR2-tdTomato (C), and Myh11CreER^T2-ChR2-tdTomato (E) with their corresponding brightfield image (B, D, F), respectively. Time-lapse brightfield images every 0.5 s starting at stimulation t = 0 for CkitCreER^T2-ChR2-tdTomato (G–J), PdgfrαCreER^TM-ChR2-tdTomato (K–N), and Myh11CreER^T2-ChR2-tdTomato (O–R). The I bar denotes the inner diameter at t = 0 over time, and white asterisks denote the contraction. Representative diameter trace for the popliteal cLV demonstrates spontaneous contractions with the dotted boxes indicating the optical stimulation event in the respective brightfield images of the time lapse images. Isolated cLVs from CkitCreER^T2-ChR2-tdTomato (S), PdgfrαCreER^TM-ChR2-tdTomato (T), and Myh11CreER^T2-ChR2-tdTomato (U) were stimulated with light pulses (red dashed lines) and the summation of contraction triggering for each genotype (V). Mean and SEM are shown, **** denotes p < 0.0001. Contraction recorded from at least six popliteal cLVs from 3 mice per genotype.

Confocal Ca²⁺ imaging of GCaMP6f expression driven by CkitCreER^T2, PdgfrαCreER^TM, and Myh11CreER^T2 over the lymphatic contraction cycle

Subcellular calcium transients are observed in many pacemaker cells. We imaged IALVs from CkitCreER^T2-GCaMP6f mice, which primarily resulted in expression of GCaMP6f in the large ovoid cells in the adventitia (Figure 9A), although we occasionally observed GCaMP6f expression in both LEC and LMCs (Figure 9A) as depicted in the maximum projection of the acquisition period (Figure 9—video 1) and the spatio-temporal maps (STMs). The aberrant expressions of GCaMP6f in cells that demonstrated the typical cobblestone morphology of LECs or the circumferential LMCs that exhibited Ca²⁺ flashes and diastolic Ca²⁺ transients (Figure 9D, E, green arrows) prior to contraction were not included in the CkitCreER^T2-GCaMP6f analysis. Of 39 CkitCreER^T2-GCaMP6f cells analyzed, only 1 CkitCreER^T2-GCaMP6f cell exhibited a spontaneous Ca²⁺ transient during the recording period (Figure 9B, C, Cell 7). However, the Ca²⁺ transient in that cell did not align temporally with the ‘Ca²⁺ flash’ of the LMC with incidental GCaMP6f expression (Figure 9C, D). Despite the lack of Ca²⁺ transients under the baseline conditions throughout the IALV contraction cycle, many CkitCreER^T2-GCaMP6f cells exhibited a robust and prolonged Ca²⁺ event in response to stimulation with the mast cell activator compound 48–80 (Figure 9F–H). Notably, the Ca²⁺ events in the ovoid cells elicited by administration of compound 48–80 did not acutely alter the LMC Ca²⁺ activity (Figure 9I, J). Similarly, the majority of PdgfrαCreER^TM-GCaMP6f expressing cells also largely lacked Ca²⁺ transients and resulted in incidental LMC GCaMP6f expression (Figure 10B, Figure 10—video 1). Some cells exhibited high basal Ca²⁺ levels (Figure 10A, D) sustained throughout the recording, but without oscillations (Figure 10B, C). In contrast, spurious GCaMP6f expression in a circumferentially oriented LMC displayed Ca²⁺ flashes associated with contraction (Figure 10B, C). Of the 21 PdgfrαCreER^TM -GCaMP6f cells assessed, only 3 exhibited Ca²⁺ transients, and those were singular events contained within a single cell within the 20-s imaging period (Figure 10E, F). The lack of either global or consistent Ca²⁺ transients within either CkitCreER^T2-GCaMP6f or PdgfrαCreER^TM-GCaMP6f IALVs was in stark contrast to Ca²⁺ imaging of Myh11CreER^T2-GCaMP6f IALVs. Myh11CreER^T2 drove GCaMP6f expression in nearly all circumferential LMCs (Figure 11A), which exhibited global and nearly synchronous Ca²⁺ flashes in 100% of the analyzed cells (Figure 11B, C). Additionally, non-synchronous stochastic and localized Ca²⁺ transients were commonly observed in the LMCs during diastole (Figure 11D, E, Figure 11—video 1). Many LMCs exhibited Ca²⁺ transients during each diastolic period while other LMCs displayed few Ca²⁺ transients or lacked diastolic Ca²⁺ transients during the recording period (Figure 11B). In aggregate, only 1 of 39 CkitCreER^T2-GCaMP6f cells and 3 of 21 PdgfrαCreER^TM-GCaMP6f cells displayed a Ca²⁺ transient during recording, while 20 of 43 LMCs displayed at least one diastolic transient apart from 43 of 43 LMCs with global Ca²⁺ flashes.

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CkitCreER^T2 drives GCaMP6f expression primarily in mast cells in mouse IALVs.

Representative max projection (A) of GCaMP6f signal over time in an IALV isolated from a CkitCreER^T2-GCaMP6f mouse with ROI indicated around individual cells, primarily large ovoid cells, but also including a circumferential lymphatic muscle cell (LMC) (cell 10) and a horizontal lymphatic endothelial cell (LEC) (cell 11). Of cells 1–9, only cell 7 had any Ca²⁺ activity (red arrows) during the recording time as indicated by the spatio-temporal maps (STMs) from each ROI (B) and their normalized F/F₀ plots in (C). In contrast, the LMC in ROI 10 had both rhythmic global Ca²⁺ events (D) that spanned the cell axis (vertical axis) in the STM (E) in addition to localized Ca²⁺ events intervening the time between global events (green arrows). Representative max projection of GCaMP6f signal over time after stimulation with C48–80 (F) with many large ovoid cells displaying long-lasting global Ca²⁺ events (G, H) while not immediately affecting the LMC Ca²⁺ dynamics (I). Calcium recordings were made in n = 6 IALVs from four mice.

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Lack of coordinated Ca²⁺ activity across contraction cycle in PDGFRα cells.

Representative max projections of GCaMP6f signal over time in IALVs isolated from PdgfrαCreER^TM-GCaMP6f mice (A, D). ROIs were made around cells and GCaMP6f recorded over time to generate the corresponding spatio-temporal maps (STMs) (B, E) for each cell and plots (C, F), respectively. Once again, incidental recombination occurred in a lymphatic muscle cell (LMC) which displayed rhythmic Ca²⁺ flashes (C) while the slight undulation in the other cells is due to movement artifact (B). Red arrows indicate the limited local Ca²⁺ activity observed in two cells from a PdgfrαCreER^TM-GCaMP6f IALV. Calcium recordings were made in n = 6 IALVs from four mice.

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Heterogeneous diastolic Ca²⁺ transient activity in lymphatic muscle cells (LMCs).

Representative max projections of GCaMP6f signal over time in an IALVs isolated from Myh11CreER^T2-GCaMP6f mice (A). LMCs were outlined with ROIs to assess GCaMp6F signal over time. Rhythmic global flashes (B) were entrained across all the LMCs in the FOV (C) with many cells exhibiting diastolic Ca²⁺ release events. Cells exhibiting at least one diastolic Ca²⁺ event, within the context of our focal plane constraints, over the recorded time were denoted by the red asterisks. The plot in (D) magnifies the first diastolic period, seconds 1–3 of C, to assist in visualizing the lack of coordination of the diastolic events. (E) Max projection of the pseudo-linescan analysis across the axis of the vessel to highlight diastolic Ca²⁺ transients in all cells in the field of view and their lack of coordination across the cells (x-axis). The white dotted box shows the first diastolic period plotted in (D). Representative images from calcium recordings from n = 4 IALVs from four mice.

Pressure dependency of subcellular Ca²⁺ transients in LMCs

We next sought to test whether diastolic Ca²⁺ transients were pressure-dependent, given that cLVs exhibit pressure-dependent chronotropy (Zawieja et al., 2019). GCaMP6f-expressing LMCs were studied at intraluminal pressures of 0.5–5 cmH₂O in the presence of nifedipine, which blocked the Ca²⁺ flashes but not local Ca²⁺ transients (Figure 12A). As intra-luminal pressure was increased, there was a marked increase in the occurrence of Ca²⁺ transients (Figure 12B, Figure 12—videos 1–3). These calcium transients were converted into particles (PTCLs) for further analysis as previously described (Drumm et al., 2019b). Activity maps of Ca²⁺ PTCL activity were generated (Figure 12C) and PTCL area (Figure 12D) and frequency were determined at each pressure (Figure 12E). The maps show that as pressure increased, the area of the LMC layer displaying a Ca²⁺ transient increased (as evident by the increase in PTCL area) as did the distribution of Ca²⁺ PTCLs across the LMC layer (Figure 12C). Across 11 experiments, the area of the field of view activated by PTCLs/frame increased from 73.2 ± 17.7 µm²/frame at 0.5 cmH₂O to 108.6 ± 20.5 µm²/frame at 2 cmH₂O and was further enhanced to 139.2 ± 26.9 µm²/frame at 5 cmH₂O (Figure 12F). The number of PTCLs per frame also increased with pressure, from 2.9 ± 0.4 at 0.5 cmH₂O to 4.1 ± 0.5 and 5.2 ± 0.6 PTCL/frame at 2 and 5 cmH₂O, respectively (Figure 12G).

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Pressure dependency of mouse lymphatic muscle cell (LMC) diastolic Ca²⁺ transients.

Representative max projection of GCaMP6f signal over 20 s in an IALVs isolated from Myh11CreER^T2-GCaMP6f mice in the presence of the L-type blocker nifedipine (1 μM) (A) and pressurized to 0.5, 2, and 5 cmH₂O. The local diastolic Ca²⁺ transients persist in the presence of nifedipine and increase with increasing pressure as demonstrated in the whole vessel spatio-temporal maps (STMs) (B). Particle occurrence maps highlight the Ca²⁺ activity in each LMC as pressure is raised (C). Representative particle analysis plots for particle area (D) and particle counts/frame at each pressure (E). Summary files for particle area (F) and count/frame (G). * denotes p < 0.05, Mean and SEM shown with n = 12 separate IALVs from 8 Myh11CreER^T2-GCaMP6f.

Contraction frequency is set by the diastolic depolarization rate

To assess how pressure regulates LMC membrane potential, we first recorded membrane potential in cells exhibiting action potentials (APs) using a microelectrode filled with biocytin-AF488 to label the impaled cell. In each case (n = 3 IALVs), the labeled cell was an LMC wrapping circumferentially around the vessel (Figure 13A, B), and as these recordings were made over the course of many minutes, the direct neighboring circumferential LMCs also exhibited fluorescence, albeit weaker in intensity, as expected for cells coupled by gap junctions (Figure 13A). In all the recorded cells exhibiting APs, we noted a diastolic depolarization preceding the sharp upstroke achieved once threshold was met at each pressure (Figure 13C). The AP frequency and rate of the diastolic depolarization increased with pressure (Figure 13D, E). Linear regression of a plot of each AP frequency and diastolic depolarization rate at each pressure demonstrated a tight association between the two parameters. However, we did not observe a significant effect of pressure on minimum membrane potential (Figure 13G), threshold potential (Figure 13H), AP upstroke (Figure 13I), AP peak potential (Figure 13J), plateau potential (Figure 13K), or the time spent over threshold (Figure 13L).

Figure 13

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Pressure-dependent diastolic depolarization in lymphatic muscle cells (LMCs).

Intracellular recordings of LMC action potentials (APs) were confirmed by loading (greater than 10 min) the impaling electrode with 1 M KCl 100 µg/ml AF488-Biocytin while recording APs followed by imaging on a spinning disk confocal microscope (n = 3 vessels from 3 mice). 3D reconstruction of the z-stack confirmed the circumferential pattern of the impaled LMC that was strongly labeled by AF488-Biocytin (A, B), which also labeled neighboring LMCs, likely through gap junctions as AF488-Biocytin is <1 kDa. In a separate set of experiments APs were recorded at three different pressures, 0.5, 2, and 5 cmH₂O. We plotted the representative recordings from one cell at each pressure (C). AP frequency was significantly increased with pressure (D) as was the diastolic depolarization rate (E). Plotting the AP frequency and diastolic depolarization rate from all recordings at each pressure (F) highlights the significant effect diastolic depolarization rate has on the AP frequency. Minimum membrane potential (G), threshold membrane potential of AP initiation (H), upstroke constant (I), peak membrane potential (J), plateau membrane potential (K), and time over threshold (L) are also reported, although not significant. Recordings are from 10 IALVs from 10 mice.

Discussion

The identification of the cellular origin and signaling mechanisms underlying cLV pacemaking will reveal novel targets for pharmacological intervention in treating lymphedema and the associated lymphatic contractile dysfunction. In this study, we tested proposed pacemaker cell types based on three parameters: (1) that the pacemaker cells are located along the entire length of the cLV, to accommodate spontaneous contractions and coordinated electrical conduction; (2) that depolarization of the pacemaker cell can drive a coordinated and propagated contraction of the vessel; and (3) that the presence of Ca²⁺ transients precedes or coincides with contraction, as commonly observed in other pacemaker cells. We used confocal microscopy and a combination of immunofluorescence and fluorescent reporters under the control of various inducible Cres to identify and target both muscle and non-muscle cells, previously labeled as ICLCs, which co-stain for the markers CD34 and PDGFRα. Our cell characterizations were supplemented by scRNAseq analysis of isolated and cleaned mouse IALVs which supported our finding of three major cell types including LECs, LMCs, and AdvCs, each of which could be further subclustered into transcriptionally unique populations. From our initial fluorescence imaging studies, a role for intrinsic pacemaking by LMCs (van Helden, 1993; von der Weid et al., 2008), or by a novel population of CD34⁺ lymphatic ICLCs (McCloskey et al., 2002; Briggs Boedtkjer et al., 2013), also referred to as telocytes, was further examined . We utilized PdgfrαCreER^TM to further test whether these cells exhibited pacemaker capabilities. However, these PDGFRα⁺ cells had minimal Ca²⁺ activity despite ongoing contractions, and optogenetic stimulation of ChR2 in these cells failed to drive a spontaneous contraction. In contrast, photo-stimulation of LMCs expressing ChR2 elicited robust, propagated contractions with similar characteristics and propagation to spontaneous contractions in the same vessels. Furthermore, Ca²⁺ imaging in LMCs revealed diastolic Ca²⁺ transients in diastole that increased in frequency and spatial spread as pressure was elevated. We also demonstrated that the primary component of the AP driving the frequency change with pressure is diastolic depolarization, which we have previously reported to be dependent on ANO1 (Zawieja et al., 2019) and IP3R1 (Zawieja et al., 2023). Notably, we recently reported that diastolic Ca²⁺ transients are abrogated in IALVs from Myh11CreER^T2-Itpr1 inducible knockout mice, supporting an IP₃R1-ANO1 axis as the pressure-dependent pacemaker mechanism in LMCs. These results, in addition to the recent findings using targeted deletion of Gjc1 (Castorena-Gonzalez et al., 2018b) or Cacna1c (To et al., 2020; Davis et al., 2022) in lymphatic muscle, support the model of LMCs as the intrinsic pacemaker, as has been previously proposed (van Helden, 1993; Helden and Crowe, 1996; van Helden and Zhao, 2000).

Pacemaking in smooth muscle

In many smooth muscle organs, regulation of a coordinated contraction is a complex and multicellular phenomenon. Multiple cell types integrate physical and biological information into electrical activity to be transmitted to the force-producing smooth muscle cells, sometimes across great distances relative to cell size, to regulate Ca²⁺ influx by voltage-dependent Ca²⁺ channels required for contraction. The intestine is one such documented tissue in which cKIT⁺ ICCs and PDGFRα⁺ interstitial cells form an electrical syncytium to regulate intestinal motility (Sanders et al., 1999; Sanders et al., 2014). The pacemaking function of intestinal ICCs relies heavily on ANO1, a Ca²⁺-activated Cl^- channel, which is required for slow wave activity in the ICCs. Both cKIT and ANO1 can be used as a marker for ICCs in the intestine (Hwang et al., 2009; Cobine et al., 2017; Malysz et al., 2017), cKIT⁺ and VIMENTIN⁺ ICLCs have been observed in sheep lymphatic vessels (McCloskey et al., 2002), yet these cell populations did not form gap junctions with the smooth muscle to form electrical connections (Briggs Boedtkjer et al., 2013) as occurs in the intestinal ICCs. Our cKIT staining and CkitCreER^T2-Rosa26^mTmG reporter studies on mouse IALVs revealed a sparse population of large ovoid cells previously classified as mast cells (Chatterjee and Gashev, 2012; Zawieja et al., 2019). Their identity as mast cells was further supported by a sustained global Ca²⁺ event after stimulation with the mast cell degranulating agent compound 48–80. However, both VIMENTIN and CD34 showed robust staining throughout the mouse lymphatic vessel wall. LECs stained for VIMENTIN, as did non-muscle stellate-shaped cells, with many co-expressing CD34. Other smaller circular cells, some of which were also cKIT⁺ as well and some whose morphology was similar to that of the macrophage staining profile of the GFP⁺ cells in IALVs from MacGreen mice, were also VIMENTIN⁺, consistent with previous reports of macrophage staining in cLVs (Bridenbaugh et al., 2013b; Chakraborty et al., 2015; Zawieja et al., 2016). While VIMENTIN⁺ cells had a perinuclear staining profile, CD34 demarcated the cell membrane and was useful for assessing the morphology of these cells. Of particular interest, the VIMENTIN⁺CD34⁺ cells densely populated the length of the mouse IALV, with a majority displaying a flattened stellate morphology characterized by a classic rounded oak-leaf appearance, although some displayed fine dendrite-like extensions. Contrasting with the previous findings in the human thoracic duct (Briggs Boedtkjer et al., 2013), we did not observe a significant population of CD34⁺ cells with a bipolar morphology oriented axially along the vessel. However, z-stack reconstructions of sections of the mouse IALV that included the secondary valves revealed interstitial CD34⁺PDGFRα⁺ cells that resembled those bipolar cells with multiple axon-like extensions throughout the endothelial leaflets; these were similar to interstitial cells that were previously reported in collecting vessel valves (Leak and Burke, 1968) and lymphovenous valves (Geng et al., 2016). While these cells have not been frequently described in the valves of peripheral cLVs, we observed them in each of the valve regions imaged and, in addition, they were labeled with other Cre drivers, including Cspg4Cre-Rosa26^mTmG and PdgfrβCreER^T2-Rosa26^mTmG (data not shown). Whether these cells regulate leaflet extracellular matrix deposition or lymphatic valve integrity is unknown, but a possible role as a critical pacemaker can be excluded as vessel segments without valves display normal contractile behavior (van Helden, 1993; Gashev et al., 2002). Instead, the majority of the CD34⁺PDGFRα⁺ cells were found in the adventitia, in two to three layers, overtop the LMCs, and they were consistently observed in high density along the IALV. Some CD34⁺PDGFRα⁺ cells or their extensions were present between the lymphatic endothelial and muscle layers, as had been previously reported with electron microscopy of human lymphatic vessels (Briggs Boedtkjer et al., 2013). Thus, while some of these AdvCs may be contained within the extracellular matrix that retracts onto the vessel during microdissection, many others are intimately dispersed within the vessel wall.

PDGFRα⁺CD34⁺ cells are not involved in cLV pacemaking under physiological conditions

Co-staining of CD34 and PDGFRα has recently been ascribed as a delineating feature of telocytes, although PDGFRα routinely labels fibroblasts and specific interstitial cells in the GI tract involved in purinergic neurotransmission in the GI tract (Kurahashi et al., 2011; Kurahashi et al., 2013; Clayton et al., 2022). Cd34 expression is also ascribed to some multipotent cell populations of various origins (Sidney et al., 2014). For example, PDGFRα⁺ fibroblasts appear to be progenitors of the smooth muscle fibers associated with the lacteal, the lymphatic capillary in the villus (Sanketi et al., 2024). It remains controversial to what extent telocytes are distinct from or are components/subtypes of either cell type and morphological discrimination between the populations typically requires electron microscopy imaging (Clayton et al., 2022). Mesenchymal stromal cells (Andrzejewska et al., 2019) and fibroblasts (Muhl et al., 2020; Buechler et al., 2021; Forte et al., 2022) are not monolithic in their expression patterns displaying both organ-directed transcriptional patterns as well as intra-organ heterogeneity (Lendahl et al., 2022) as readily demonstrated by recent single-cell RNA sequencing studies that provided immense detail about the subtypes and activation spectrum within these cells and their plasticity (Luo et al., 2022b). We were able to distinguish up to 10 subclusters of AdvCs, the majority of which expressed or co-expressed Cd34 and Pdgfra. These cells were consistently negative for smooth muscle markers such as Des, Cnn1, Acta2, Myh11, or the pericyte marker Mcam. However, Pgfrb expression was noted in our scRNAseq analysis and in our RT-PCR of sorted PdgfrαCreER^TM-Rosa26^mTmG cells. PDGFRβ protein expression was confirmed with variable immunofluorescence staining amongst the PDGFRα stained cells as well as LMCs. The PdgfrβCreER^T2Rosa26^mTmG mice had only modest recombination in both the LMC and PDGFRα⁺ cell populations, but potentially highlighted a myofibroblast-like cell subpopulation, cells that might lie on the spectrum of differentiation from lymphatic muscle and PDGFRα⁺ cells, or perhaps a cell with pacemaker activity as PDGFRβ is widely used as a pericyte marker and some pericytes display pacemaker activity (Hashitani et al., 2015). Adding to this intrigue, the PdgfrαCreER^TM sorted cells expressed transcripts for Cacna1c, the voltage-gated L-type Ca²⁺ channel critical for lymphatic contractions (Zawieja et al., 2018a; To et al., 2020); Ano1, the ion channel underlying pressure-dependent chronotropy (Mohanakumar et al., 2018; Zawieja et al., 2019); and Gjc1, the primary connexin mediating electrical conduction in mouse lymphatic collecting vessels (Castorena-Gonzalez et al., 2018b; Hald et al., 2018). Expression of these genes in certain subpopulations of the AdvCs was also apparent in our scRNAseq analysis. Thus, the presence of those gene transcripts does not appear to be due to muscle cell contamination or incidental recombination in LMCs, as we did not detect LMC markers in the RT-PCR profiling of the sorted PDGFRα⁺ cells, nor were GFP-expressing cells with an LMC morphology observed in imaging of PdgfrαCreER^TM-Rosa26^mTmG vessels. Critically, however, deletion of Cacna1c, Gjc1, or Ano1 through PdgfrαCreER^TM-mediated recombination neither recapitulated the previous phenotypes achieved with Myh11CreER^T2 (Castorena-Gonzalez et al., 2018b; Zawieja et al., 2019; To et al., 2020; Davis et al., 2022) nor significantly affected pacemaking in mouse popliteal cLVs. This finding is in stark contrast to the complete lack of contractions observed in Myh11CreER^T2-Cacna1c ^fl/fl vessels (To et al., 2020; Davis et al., 2023b) or the vessels from vascular muscle specific Itga8CreER^T2-Cacna1c^fl/fl mice (Davis et al., 2022; Warthi et al., 2023), and the significant loss in pressure-induced chronotropic modulation of pacemaker function in IALVs with Myh11CreER^T2-mediated deletion of Ano1 that we have previously reported (Zawieja et al., 2019). While a subpopulation of CD34⁺PDGFRα⁺ cells may share expression of critical pacemaker genes identified in the LMCs, they do not appear to be involved in cLV pacemaking or contractile function under physiological states. Instead, CD34⁺PDGFRα⁺ cells co-stained significantly with LY6A⁺, suggesting they may be primed to act as resident multipotent cells (Song et al., 2020; Kimura et al., 2021). To this point, the PdgfrαCreER^TM FACS-purified cells also expressed markers associated with ‘stemness’such as Cd34, Klf4, Gli1, Itgb1, Eng, Cd44, and Vimentin, in addition to Ly6a, and it is likely that the PdgfrαCreER^TM population includes various distinct subpopulations (Jolly et al., 2022) expressing these markers. These cells may play a role in rebuilding the lymphatic collecting vessel vasculature following collecting vessel damage or lymph node resection, and further studies are required to assess their functional multipotency.

SR Ca²⁺ cycling in pacemaking

The use of the mouse IALV model, in addition to the simplicity of the vessel architecture, provided the use of genetic tools that previously had been instrumental in identifying the cKIT⁺ ICC as the pacemaker cells of the GI tract (Ward et al., 1994; Huizinga et al., 1995; Torihashi et al., 1995). Through the use of the respective PdgfrαCreER^TM and Myh11CreER^T2 drivers, we were able to specifically image Ca²⁺ in each respective cell type in pressurized, contracting vessels. Pacemaking initiating cells have an inherently unstable membrane potential, oftentimes utilizing the oscillatory nature of Ca²⁺ release from the sarcoendoplasmic reticulum coupled to Ca²⁺-sensitive electrogenic exchangers or ion channels to drive depolarization (van Helden, 1993; Hashitani et al., 2015; Baker et al., 2021b; Sanders et al., 2022). One such example is the pacemaker ICC in the gastric corpus which exhibits abundant Ca²⁺ transients that couple to ANO1-mediated chloride currents in both the intervening period between slow waves as well as the plateau phase of the slow wave (Baker et al., 2021a); however, such activity is not characteristic of all pacemaker ICC types. The identification of a Ca²⁺-activated chloride current in LMCs (van Helden, 1993; Toland et al., 2000) and its correspondence with subcellular Ca²⁺ transients (van Helden, 1993; Ferrusi et al., 2004; von der Weid et al., 2008) led Van Helden to postulate that LMCs had intrinsic pacemaking capability (van Helden, 1993; Helden and Crowe, 1996). We have previously reported that mouse LMCs in pressurized vessels routinely display subcellular Ca²⁺ release events that reflect the kinetics and characteristics of Ca²⁺ puffs and waves in addition to the coordinated global Ca²⁺ flash associated with Ca²⁺ influx during an AP (Castorena-Gonzalez et al., 2018b; Zawieja et al., 2018a; Zawieja et al., 2019). Here we confirmed the consistent presence of subcellular Ca²⁺ transients only in LMCs with GCaMP6f driven by Myh11CreER^T2 but not in the cells with GCaMP6f driven by PdgfrαCreER^TM. Critically, we also demonstrated that the Ca²⁺ transients increased in both frequency and spatial spread as pressure was elevated in the vessel, as would be expected to account for the pressure-dependent chronotropy observed in lymphatic collecting vessels. This underscores the finding that genetic deletion of Ano1 in the LMCs dramatically reduced contraction frequency and abolished pressure-dependent chronotropy in those vessels (Zawieja et al., 2019). This phenotype was largely replicated with a similar reduction in frequency and loss of pressure-dependent chronotropy in our recent study utilizing Myh11CreER^T2 to drive deletion of IP3R1 from LMCs (Zawieja et al., 2023) in which these diastolic Ca²⁺ transients were absent. This fits with the central role of IP3R and subcellular Ca²⁺ release as critical components of intrinsic LMC pacemaking (Helden and Crowe, 1996; von der Weid et al., 2008). In addition to the transcriptional heterogeneity identified by scRNASeq, we also noted heterogeneity in the propensity of LMCs to display diastolic Ca²⁺ transients under control conditions or show the sustained Ca²⁺ oscillations that occur in the presence of nifedipine. We did not detect any significant difference in the expression of Itpr1, the gene encoding the IP3R1, across our LMCs subclusters. However, when using less stringent conditions, we identified that the LMC cluster “0” had significantly increased expression of Itprid2 (Log2FC of 0.26), which encodes the Kras-induced actin-interacting protein (KRAP). KRAP has recently been implicated in IP3R1 immobilization and licensing and was required for IP3R1-mediated Ca²⁺ puffs (Thillaiappan et al., 2021; Atakpa-Adaji et al., 2024). Whether the higher expression of KRAP results in a greater probability of these LMCs to display IP3R1-dependent Ca²⁺ oscillations in LMCs requires further investigation. Of note, LMCs also express the components for store-operated calcium entry including Stim1, Stim2, Orai1, Orai3, Saraf, and Stimate, which may be involved in maintaining IP3R1-dependent SR Ca²⁺ release oscillations.

The membrane potential recordings made in this study suggest that the regulation of pressure-dependent chronotropy is through modulation of the diastolic depolarization rate in LMCs, as previously demonstrated in rat mesenteric lymphatic vessels (Zawieja et al., 2018b). The appearance of the diastolic depolarization may depend on the method of vessel stretch employed as it is not always observed in wire myograph preparations (von der Weid et al., 2014). Notably, in this study, PdgfrαCreER^TM-mediated deletion of Ano1 had no effect on contractile parameters. The lack of Ca²⁺ transients in PDGFRα⁺ cells across any stage of the lymphatic contraction cycle also diminishes any expected role for this cell type to perform as the pacemaker for the mouse IALV. We recently showed that pressure-dependent Ca²⁺ mobilization from the SR, through IP3R1 (Zawieja et al., 2023), sets the basis for LMC pacemaking as previously proposed (van Helden, 1991; von der Weid et al., 2008). However, the mechanisms driving IP3R1 activation and Ca²⁺ oscillations remain to be fully addressed.

A pacemaker cell would be expected to be electrically coupled to the LMC layer to permit the nearly synchronous conduction velocity of the contraction wave (Zawieja et al., 1993; Castorena-Gonzalez et al., 2018b; Hald et al., 2018) and to transmit depolarization into coupled LMCs to activate the voltage-dependent Ca²⁺ channels that are responsible for lymphatic muscle APs. Connexins are the molecular constituents of gap junctions and, as stated above, we detected Gjc1 expression in PdgfrαCreER^TM sorted cells. However, we did not detect any impairment in pacemaking, nor were contraction conduction speed deficits or multiple pacemakers noted in the PdgfrαCreER^TM -Gjc1^fl/fl popliteal cLVs, in contrast to the development of multiple pacemaker sites and the lack of entrainment that characterize cLVs from Myh11CreER^T2-Gjc1^fl/fl mice (Castorena-Gonzalez et al., 2018b). Admittedly, we did not perform an exhaustive assessment of the connexin expression profile of the CD34⁺PDGFRα⁺ cells, and Gjc1 may not be the dominant connexin expressed in the CD34⁺PDGFRα⁺ cells, or heterotypic connexons could exist (Koval et al., 2014). However, electron microscopy studies of the putative ICLC in the human thoracic duct did not detect any gap junctions, although peg-and-socket connections were observed (Briggs Boedtkjer et al., 2013). We utilized optogenetics to directly depolarize the specific cell populations in both the PdgfrαCreER^TM and Myh11CreER^T2 mouse models in an attempt to drive contractions. Local photo-stimulation of the PDGFRα cells failed to initiate contraction, while the stimulation of Myh11CreER^T2 recombined cells resulted in contractions that were indistinguishable from the spontaneously occurring contractions. These results give functional credence to the lack of hetero-cellular coupling via gap junctions that was previously reported (Briggs Boedtkjer et al., 2013). Just as critically, our results also highlight the regenerative nature of the lymphatic muscle AP. Local, optogenetic-initiated depolarization of either a single or a few LMCs to threshold was sufficient to drive a coordinated contraction along the vessel demonstrating conducted activity at the tissue level.

Conclusions

Our present findings lend further support to the hypothesis that the LMCs are intrinsic pacemakers (van Helden et al., 2006; Mitsui and Hashitani, 2020) and that mouse cLVs do not require an ICC-like cell network to drive propagated contractions. These findings also underscore the significance of lymphatic muscle Ca²⁺ handling as the driver of lymphatic pacemaking, which can be compromised in disease states leading to impaired lymphatic contractile activity (Stolarz et al., 2019; Lee et al., 2020; Van et al., 2021). Further studies delineating the specific SR Ca²⁺ release and influx pathways, and the contributions of Ca²⁺ sensitive ion channels are required to develop sophisticated in silico models and identify potential therapeutic targets to rescue lymphatic pacemaking in lymphedema patients (Olszewski, 2002; Olszewski, 2008).

Limitations

One fundamental assumption underlying our conclusions is that there is a conserved pacemaking pathway and cell type regulating lymphatic collecting vessel contractions across species, specifically pertaining to the capability of lymphatic muscle to maintain pacemaking and coordination despite changes in tissue complexity and cLV wall thickness. It is worth noting that lymphatic collecting vessels in mice have similar pressure-dependent chronotropy and contraction conduction velocity as recorded in rats and human vessels (Castorena-Gonzalez et al., 2018b). These similarities exist despite the fact that mouse lymphatic collecting vessels are typically encircled by a single layer of lymphatic muscle, while larger species may have multiple layers of LMCs in the wall. It is possible that vessels with multiple layers of LMCs need a network of ICLC to coordinate their activity. The simplicity in the makeup of the mouse cLV and the use of cell targeting Cre models provides great control over experimental variables, but other cell types may be required for coordination of LMC pacemaking in other species where the lymphatic cLV walls are larger and thicker and contain multiple muscle cell layers. Our scRNAseq analysis also is likely biased using Rosa26^mTmG mice with FACS purification to remove debris and concentrate specific cell types from these pooled small vessels. Larger and more complex cells, with attributes that can be ascribed to ICCs, are more likely to be lost in this methodology (e.g., depending on the FACS gating parameters), and this procedure can also elicit a stress response in the transcriptome of the analyzed cells. However, we also did not observe long and complex cells, aside from the circumferential LMCs, in our immunofluorescence and recombination reporter imaging experiments. Immediate and early gene expression motifs driven by a stress response may be a component of the differences in subclusters that were identified. Future scRNAseq or snRNAseq studies with deeper sequencing will be required to ensure that the full transcriptomic heterogeneity is accounted for under different cellular stress conditions.

Our data demonstrate that limited staining of a few cell markers alone is insufficient to identify discrete cell populations in mouse cLVs. Additionally, mRNA expression does not equal protein translation nor guarantee specific function as we did not readily detect endothelial CD34 with immunofluorescence despite detecting transcript; additionally, PdgfrαCreER^TM-mediated deletion of Ano1, Gjc1, or Cacna1c had no effect on cLV pacemaking. Further experimentation is also required to fully characterize expression of multipotent cell markers and function of CD34⁺PDGFRα⁺LY6A⁺ cells invested within the mouse cLVs, although doing so was beyond the scope of this study assessing pacemaker identity. Tangentially, another limitation of our approach pertains to the specificity and recombination efficiency of inducible Cre recombinase models, which can be a notable confounding variable (Chakraborty et al., 2019). We observed that our inducible Cre models led to a degree of nonspecific recombination within the mouse cLV, with GCaMP6f and ChR2 particularly susceptible to recombination compared to the Rosa26^mTmG reporter. Recombination in multiple cell types was expected with the constitutive Cre models we employed (Cspg4Cre and PdgfrαCre), as vascular and lymphatic muscle precursor cells can transiently express Nestin, Pdgfra, and Cspg4 (Hill et al., 2015; Castorena-Gonzalez et al., 2018b; Kenney et al., 2020). We also observed that PdgfrβCreER^T2 drove recombination in a subpopulation of LMCs and PDGFRα⁺ cells. These appeared to be two distinct populations that only share expression for Pdgfrb based on our scRNAseq dataset, but which may exist along a continuum of differentiation. PDGFB–PDGFRβ signaling is critical for normal mural cell recruitment to both the blood and lymphatic vasculature (Gaengel et al., 2009; Wang et al., 2017) and proliferating vascular smooth muscle cells and pericytes have both been documented to express Pdgfrb (Andrae et al., 2008; Pitulescu and Adams, 2014). Ideally, novel Cre or combinatorial Cre models that specifically target LMCs or subpopulations of LMCs may be developed to further tease out the functional roles of these cells.

Materials and methods

Mice

Wild-type male mice (25–35 g) on the C57BL/6J background, Rosa26^mTmG reporter (Muzumdar et al., 2007) (Strain#007676), transgenic PdgfrαCre (Strain#013148), CSFR1-EGFP (MacGreen) (Sasmono et al., 2003) (Strain#018549), genetically encoded Ca²⁺ sensor GCaMP6f (Chen et al., 2013) (Strain#028865), transgenic PdgfrαCreER^TM (Kang et al., 2010) (Strain#018280), Cspg4-Cre (Strain #:008533) (Zhu et al., 2008), and ChR2/tdTomato fusion mice (Madisen et al., 2012) (Strain#012567) were purchased from The Jackson Laboratory (Bar Harbor, MA, USA). PdgfrβCreER^T2 (Gerl et al., 2015) mice were a gift from Ralf Adams (Mac Planck Institute) and kindly provided by Lorin Olson (Oklahoma Medical Research Foundation) and are currently available at Jax (Strain#029684). The Myh11CreER^T2 mice (Wirth et al., 2008) were a gift from Stefan Offermanns, Max-Planck-Intstitut für Herz- und Lungendforschung, Bad Nauheim, Germany, and are currently available at Jax (Strain #019079, Y-Linked). CkitCreER^T2 mice (Heger et al., 2014) were a gift from Dieter Saur (Technical University of Munich). Prox1-eGFP mice (Choi et al., 2011) were a gift from Young-Kwon Hong (University of Southern California). For genotyping, we isolated genomic DNA from mouse tail clips using the HotSHOT method (Truett et al., 2000). Specific mouse genotypes were confirmed via PCR using 2x PCR Super Master Polymerase Mix (Catalog # B46019, Bimake, Houston, TX) performed as specified by the provider. Mice used for this study were 3–12 months of age. All animal protocols were approved by the University of Missouri Animal Care and Use Committee (Protocol 53461 and 41500) and conformed to the US Public Health Service policy for the humane care and use of laboratory animals (PHS Policy, 1996).

Gene	Strand	Accession #	Sequence (5'–3')	Size	Exon	Source
Prox1	s	NM_008937	GTA AGA CAT CAC CGC GTG C	218	1	NIH Primer Tool
Prox1	as		TCA TGG TCA GGC ATC ACT GG		2
Itgam (Cd11b)	s	NM_008401	ATG GAC GCT GAT GGC AAT ACC	203	13	MGH Primer Bank ID 668048a1
Itgam (Cd11b)	as		TCC CCA TTC ACG TCT CCC A		14
Pdgfra	s	NM_011058	AGA GTT ACA CGT TTG AGC TGT C	252	8	MGH Primer Bank 26349287a1
Pdgfra	as		GTC CCT CCA CGG TAC TCC T		10
Myh11	s	NM_013607	AAG CTG CGG CTA GAG GTC A	238	33	MGH Primer Bank ID 7305295a1
Myh11	as		CCC TCC CTT TGA TGG CTG AG		34
cKit (Cd117)	s	NM_021099	CGC CTG CCG AAA TGT ATG ACG	162	21	Drumm et al., 2018
cKit (Cd117)	as		GGT TCT CTG GGT TGG GGT TGC		23
Pdgfrb	s	NM_008809	AGC TAC ATG GCC CCT TAT GA	367	16	Basciani et al., 2004
Pdgfrb	as		GGA TCC CAA AAG ACC AGA CA		19
Cdh5 (Cadherin, VE-cadherin)	s	NM_009868	CTT CCT TAC TGC CCT CAT TGT	313	3	IDT Primer Quest
Cdh5 (Cadherin, VE-cadherin)	as		CTG TTT CTC TCG GTC CAA GTT		5
Nos3 (eNOS)	s	NM_008713	CTG CCA CCT GAT CCT AAC TTG	143	22	IDT Real time primer tool
Nos3 (eNOS)	as		CAG CCA AAC ACC AAA GTC ATG		23
Acta2 (Smooth Muscle Actin)	s	NM_007392	GAG CTA CGA ACT GCC TGA C	129	7	IDT TaqMan Mm.PT.58.16320644
Acta2 (Smooth Muscle Actin)	as		CTG TTA TAG GTG GTT TCG TGG A		8
Cacna1c exon1b	s	NM_001159533	ATG GTC AAT GAA AAC ACG AGG ATG		1	Cheng et al., 2007
Cacna1c exon1b	as		GGA ACT GAC GGT AGA GAT GGT TGC	234	2
Cd34	as	NM_133654	GGT ACA GGA GAA TGC AGG TC	119	1	IDT Mm.PT.58.8626728
Cd34	s		CGT GGT AGC AGA AGT CAA GT		2
Cspg4 (Ng2)	as	NM_139001	CTT CAC GAT CAC CAT CCT TCC	132	5	IDT Mm.PT.58.29461721
Cspg4 (Ng2)	s		CCC GAA TCA TTG TCT GTT CCC		6
Vimentin	s	NM_011701	CTG TAC GAG GAG GAG ATG CG	249	1	Li et al., 2016
Vimentin	as		AAT TTC TTC CTG CAA GGA TT		3
Desmin	s	NM_010043	GTG GAT GCA GCC ACT CTA GC	218	3	MGH Primer Bank ID 33563250a1
Desmin	as		TTA GCC GCG ATG GTC TCA TAC		4
Mcam (Cd146)	s	NM_023061	CCC AAA CTG GTG TGC GTC TT	220	1	MGH Primer Bank 10566955a1
Mcam (Cd146)	as		GGA AAA TCA GTA TCT GCC TCT CC		3
Klf4	s	NM_010637	ATT AAT GAG GCA GCC ACC TG	400	1	Majesky et al., 2017
Klf4	as		GGA AGA CGA GGA TGA AGC TG		3
Ly6a (Sca1)	s	NM_001271416	CTC TGA GGA TGG ACA CTT CT	400	2	Majesky et al., 2017
Ly6a (Sca1)	as		GGT CTG CAG GAG GAC TGA GC		4
Gli1	s	NM_01029	ATC ACC TGT TGG GGA TGC TGG AT	316	8	Kramann et al., 2015
Gli1	as		CGT GAA TAG GAC TTC CGA CAG		10
Itgb1 (Cd29)	s	NM_010578	TCG ATC CTG TGA CCC ATT GC	170	14	NIH Primer Tool
Itgb1 (Cd29)	as		AAC AAT TCC AGC AAC CAC GC		15
Endoglin (Eng, Cd105)	s	NM_007932	TGA GCG TGT CTC CAT TGA CC	416	11	NIH Primer Tool
Endoglin (Eng, Cd105)	as		GGG GCC ACG TGT GTG AGA A		15

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Strain, strain background (Mus musculus)	C57BL/6J	The Jackson Laboratory	Jax Strain #000664 RRID:IMSR_JAX:000664
Strain, strain background (Mus musculus)	Rosa26^mTmG	The Jackson Laboratory	Jax Strain #007676 B6.129(Cg)-Gt(ROSA)26Sortm4(ACTB-tdTomato,-EGFP) Luo/J RRID:IMSR_JAX:007676
Strain, strain background (Mus musculus)	PdgfrαCre	The Jackson Laboratory	Jax Strain #013148 C57BL/6-Tg(Pdgfra-cre)1Clc/J RRID:IMSR_JAX: 013148
Strain, strain background (Mus musculus)	Csfr1-EGFP	The Jackson Laboratory	Jax Strain #018549 B6.Cg-Tg(Csf1r-EGFP)1Hume/J RRID:IMSR_JAX:018549
Strain, strain background (Mus musculus)	PdgfrαCreER^TM	The Jackson Laboratory	Jax Strain #018280 B6N.Cg-Tg(Pdgfra-cre/ERT)467Dbe/J RRID:IMSR_JAX:018280
Strain, strain background (Mus musculus)	Cspg4-Cre	The Jackson Laboratory	Jax Strain #:008533 B6;FVB-Ifi208Tg(Cspg4-cre)1Akik/J RRID:IMSR_JAX:008533
Strain, strain background (Mus musculus)	ChR2/tdTomato	The Jackson Laboratory	Jax Strain #012567 B6.Cg-Gt(ROSA)26Sortm27.1(CAG-COP4*H134R/tdTomato)Hze/J RRID:IMSR_JAX:012567
Strain, strain background (Mus musculus)	PdgfrβCreER^T2	The Jackson Laboratory	Jax Strain #029684 B6.Cg-Tg(Pdgfrb-cre/ERT2)6096Rha/J RRID:IMSR_JAX:029684
Strain, strain background (Mus musculus)	Myh11CreER^T2	The Jackson Laboratory	Jax Strain #019079 B6.FVB-Tg(Myh11-icre/ERT2)1Soff/J RRID:IMSR_JAX:019079
Strain, strain background (Mus musculus)	CkitCreER^T2	Dieter Saur (Technical University of Munich)	Kittm1(cre/ERT2)Dsa
Strain, strain background (Mus musculus)	Prox1-eGFP	Young-Kwon Hong (University of Southern California)	MMRRC ID #31006 Tg(Prox1-EGFP)KY221Gsat/Mmucd RRID:MMRRC_031006-UCD
Strain, strain background (Mus musculus)	GCaMP6f	Jax	Jax Strain #028865 Ai95(RCL-GCaMP6f)-D (C57BL/6J) or Ai95D (C57BL/6J) RRID:IMSR_JAX:028865
Antibody	anti-ACTA2	Sigma-Aldrich	Cat# A2547, RRID:AB_476701	(IF) 1:500
Antibody	anti-GFP	Thermo Fisher Scientific	Cat# A-11122, RRID:AB_221569	(IF) 1:200
Antibody	anti-CKIT	Cell Signaling	Cell Signaling Technology Cat# 3074, RRID:AB_1147633	(IF) 1:100
Antibody	anti-VIMENTIN	Thermo Fisher Scientific	Cat# OMA1-06001, RRID:AB_325529	(IF) 1:100
Antibody	anti-DESMIN	Invitrogen	Cat# PA5-16705, RRID:AB_10977258	(IF) 1:200
Antibody	anti-GFP	Abcam	Cat# ab13970, RRID:AB_300798	(IF) 1:200
Antibody	anti-CD34 (RAM34)	Thermo Fisher Scientific	Cat# 14-0341-82, RRID:AB_467210	(IF) 1:200
Antibody	anti-PDGFRα	R&D Systems	Cat# AF1062, RRID:AB_2236897	(IF) 1:200
Antibody	anti-PDGFRβ	eBiosciences	Cat# 14-1402-82, RRID:AB_467493	(IF) 1:200
Antibody	anti-CALPONIN	Abcam	Cat# ab46794, RRID:AB_2291941	(IF) 1:500
Antibody	anti-LY6A	Biolegend	Cat# 108101, RRID:AB_313338	(IF) 1:200
Sequence-based reagent	Prox1- Forward NM_008937	NIH Primer Tool, this paper	PCR primers	GTA AGA CAT CAC CGC GTG C
Sequence-based reagent	Prox1- Reverse NM_008937	NIH Primer Tool, this paper	PCR primers	TCA TGG TCA GGC ATC ACT GG
Sequence-based reagent	Itgam Reverse NM_008401	MGH Primer Bank ID 668048a1	PCR primers	ATG GAC GCT GAT GGC AAT ACC
Sequence-based reagent	Itgam Forward NM_008401	MGH Primer Bank ID 668048a1	PCR primers	TCC CCA TTC ACG TCT CCC A
Sequence-based reagent	Pdgfra Forward NM_011058	MGH Primer Bank ID 26349287a1	PCR primers	AGA GTT ACA CGT TTG AGC TGT C
Sequence-based reagent	Pdgfra Reverse NM_011058	MGH Primer Bank ID 26349287a1	PCR primers	GTC CCT CCA CGG TAC TCC T
Sequence-based reagent	Myh11 Forward NM_013607	MGH Primer Bank ID 7305295a1	PCR primers	AAG CTG CGG CTA GAG GTC A
Sequence-based reagent	Myh11 Reverse NM_013607	MGH Primer Bank ID 7305295a1	PCR primers	CCC TCC CTT TGA TGG CTG AG
Sequence-based reagent	Ckit Forward NM_021099	Drumm et al., 2018	PCR primers	CGC CTG CCG AAA TGT ATG ACG
Sequence-based reagent	Ckit Reverse NM_021099	Drumm et al., 2018	PCR primers	GGT TCT CTG GGT TGG GGT TGC
Sequence-based reagent	Pdgfrb Forward NM_008809	Basciani et al., 2004	PCR primers	AGC TAC ATG GCC CCT TAT GA
Sequence-based reagent	Pdgfrb Reverse NM_008809	Basciani et al., 2004	PCR primers	GGA TCC CAA AAG ACC AGA CA
Sequence-based reagent	Cdh5 Forward NM_009868	IDT Primer Quest Tool, this paper	PCR primers	CTT CCT TAC TGC CCT CAT TGT
Sequence-based reagent	Cdh5 Reverse NM_009868	IDT Real time primer tool, this paper	PCR primers	CTG TTT CTC TCG GTC CAA GTT
Sequence-based reagent	Nos3 Forward NM_008713	IDT Real time primer tool, this paper	PCR primers	CTG CCA CCT GAT CCT AAC TTG
Sequence-based reagent	Nos3 Reverse NM_008713	IDT Real time primer tool, this paper	PCR primers	CAG CCA AAC ACC AAA GTC ATG
Sequence-based reagent	Acta2 (Smooth Muscle Actin) Forward NM_007392	IDT TaqMan Mm.PT.58.16320644	PCR primers	GAG CTA CGA ACT GCC TGA C

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Methylene blue staining of isolated mouse IALVs.

Staining mouse IALVs for ICLC markers.

Immunofluorescence labeling of mouse IALVs with markers for ICLC, lymphatic muscle cell (LMC), lymphatic endothelial cell (LEC), and immune cell populations.

Inducible Cre-Rosa26mTmG labeling and fidelity to target putative pacemaker cell populations.

Figure 4—source data 1

Figure 4—source data 2

scRNAseq analysis of mouse IALVs from Rosa26mTmG mice.

RT-PCR Profiling of FACS-purified cells from inducible Cre-Rosa26mTmG.

Figure 6—source data 1

Figure 6—source data 2

Isobaric contractile assessment of popliteal collecting lymphatic vessel (cLV) from PdgfrαCreERTM-driven deletion of Ano1, GJC1, and Cacna1c.

ChR2-mediated depolarization only in lymphatic muscle cells (LMCs) triggers contraction.

CkitCreERT2 drives GCaMP6f expression primarily in mast cells in mouse IALVs.

Lack of coordinated Ca2+ activity across contraction cycle in PDGFRα cells.

Heterogeneous diastolic Ca2+ transient activity in lymphatic muscle cells (LMCs).

Pressure dependency of mouse lymphatic muscle cell (LMC) diastolic Ca2+ transients.

Pressure-dependent diastolic depolarization in lymphatic muscle cells (LMCs).

Primer list for RT-PCR.

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Scott D Zawieja

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Grace A Pea

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Sarah E Broyhill

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Advaya Patro

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Karen H Bromert

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Charles E Norton

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Hae Jin Kim

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Sathesh Kumar Sivasankaran

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Min Li

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Jorge A Castorena-Gonzalez

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Bernard T Drumm

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Michael J Davis

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Inducible Cre-Rosa26^mTmG labeling and fidelity to target putative pacemaker cell populations.

scRNAseq analysis of mouse IALVs from Rosa26^mTmG mice.

RT-PCR Profiling of FACS-purified cells from inducible Cre-Rosa26^mTmG.

Isobaric contractile assessment of popliteal collecting lymphatic vessel (cLV) from PdgfrαCreER^TM-driven deletion of Ano1, GJC1, and Cacna1c.

CkitCreER^T2 drives GCaMP6f expression primarily in mast cells in mouse IALVs.

Lack of coordinated Ca²⁺ activity across contraction cycle in PDGFRα cells.

Heterogeneous diastolic Ca²⁺ transient activity in lymphatic muscle cells (LMCs).

Pressure dependency of mouse lymphatic muscle cell (LMC) diastolic Ca²⁺ transients.