R17 Drives Heparan Sulfate Clearance via Endolysosomal Trafficking

Abstract

Heparan sulfate (HS) proteoglycans are information‐rich macromolecules that can orchestrate extracellular signaling and cargo uptake across diverse cellular contexts. Ligands that can engage cell surface HS possess significant potential as probes to investigate or manipulate cell–matrix interactions. Here, we show that R17, an HS‐binding protein from the rodent herpesvirus Peru, actively remodels the glycocalyx of pancreatic cancer cells. Recombinant R17 bound heparin in vitro and associated with the surface of pancreatic cancer cells to promote the dose‐dependent clearance of HS via trafficking to lysosomes. R17 also reduced wound closure without detectable cytotoxicity, indicative of the ability of R17 to suppress cellular migration. Notably, HS depletion persisted upon cation‐independent mannose‐6‐phosphate receptor knockdown, suggesting that clearance is independent of this mechanism. These findings identify R17 as an exogenous HS‐binding protein that can drive HS clearance via endolysosomal trafficking and suggest an alternative approach to modulate HS‐dependent functions.

R17 is a viral protein that binds heparan sulfate glycosaminoglycans and traffics them to the lysosome, leading to their degradation in a manner that does not require the presence of the cation‐independent mannose‐6‐phosphatre receptor. In BxPC3 pancreatic cancer cells, this loss in heparan sulfate results in the reduction of cell migration.

1. Introduction

Heparan sulfate glycosaminoglycans (HS GAGs) are essential mediators of cell–cell and cell–matrix communication,^{[ 1} , ² , ^{3 ]} integrating biochemical and mechanical cues across diverse biological contexts. HS binds a wide array of ligands, including growth factors, morphogens, and extracellular enzymes. Through these interactions, HS can modulate signaling pathways, morphogen gradients, and cellular uptake processes.^{[ 4} , ⁵ , ⁶ , ⁷ , ⁸ , ^{9 ]} In pancreatic cancers, HS‐related pathways are heavily implicated in disease progression.^{[ 10} , ¹¹ , ^{12 ]} Because of the important roles HS can play in cell biology, there is significant interest in identifying HS ligands across the natural and synthetic world.^{[ 13} , ^{14 ]}

In this study, we evaluate the binding characteristics and cellular effects of R17 (Figure  1A), a protein from rodent herpesvirus Peru. R17 is a potent chemokine inhibitor that also binds anionic GAGs using two basic surface clusters.^{[ 15} , ^{16 ]} Using a combination of in vitro biochemical and cell‐based approaches, we further refine this understanding and show that R17 binds to HS GAG chains. We further show that binding induces the internalization and lysosomal trafficking of R17 and HS complexes, which leads to the loss of cell surface and total HS levels. This R17‐driven process reshapes the glycocalyx and reduces the ability of BxPC3 cancer cells to migrate.

Figure 1.

R17 binds heparin in vitro. A) X‐Ray crystal structure of R17¹⁶ (PDB: 4ZKQ), highlighting basic surface clusters implicated in GAG binding. B) Heparin ELISA showing dose‐dependent binding of R17^WT (EC₅₀ 31 nM). C) Competition ELISA. Immobilized heparin plates were pre‐incubated with surfen (9 μM) followed by R17 titration. D) Competition ELISA. Immobilized heparin plates were pre‐incubated with R17 (33 nM) followed by surfen titration. Data are mean ± SD with n ≥ 2 independent experiments.

Our findings reveal that R17 acts not merely as a binding partner but as an active effector that promotes HS clearance through endolysosomal trafficking. Our work builds on the precedence that binding to cell surface HS is a common mode of viral entry.^{[ 17} , ¹⁸ , ^{19 ]} We demonstrate that in addition to acting as an HS binder, it can target HS to lysosomes for degradation.^{[ 20} , ²¹ , ²² , ²³ , ^{24 ]} By elucidating this mechanism, we further uncover an intriguing dimension of HS regulation—one in which an exogenous HS‐binding protein can directly remodel the cell surface glycome.^{[ 22 ]} These insights expand the conceptual framework of HS biology and provide new opportunities for chemical modulation of HS‐dependent pathways in both physiological and pathological contexts.

2. Results

2.1. R17 Binds Heparan Sulfate Glycosaminoglycans in Vitro

To provide further evidence that R17 (R17^WT) binds GAGs (Figure 1A) and evaluate whether HS GAGs contribute to binding, we expressed recombinant R17 in Expi293F cells with a poly‐histidine (His) tag (Figure S1, Supporting Information). Using an in vitro ELISA plate with immobilized heparin, a variant of HS that is derived from porcine intestinal mucosa, we observed dose‐dependent increases in heparin‐bound R17 (Figure 1B), corresponding with an EC₅₀ of 31 nM. An R17 mutant, where the GAG binding site is mutated (R17^GAG2) did not show significant binding to heparin in the concentrations tested (Figure S2, Supporting Information). To further ascertain that R17 binds the same sites on heparin as a known small molecule HS antagonist,^{[ 14} , ^{25 ]} we performed heparin ELISA experiments in the presence of surfen. In conditions where the heparin‐immobilized plate was pre‐incubated with surfen (9 μM) and titrated with increasing concentrations of R17, we observed a modest reduction in the potency of R17 binding, reflected by an increase in the EC₅₀ value of R17 binding to 40 nM (Figure 1C). When the heparin plate was pretreated with R17 (31 nM) and then co‐incubated with surfen at increasing concentrations, we observed that surfen was able to inhibit R17 binding (IC₅₀: 9 μM; Figure 1D).

2.2. R17 Binds HS in Cells and Causes a Dose‐Dependent Loss

Next, we sought to evaluate whether R17 can bind HS in pancreatic cancer cells. We used the BxPC3 cell line for our studies, upon determining that it expressed abundant amounts of HS (Figure S3, Supporting Information). Using flow cytometry‐based experiments, we analyzed the binding of R17 and the GAG‐mutated construct (R17^GAG2) to the surfaces of fixed BxPC3 cells and observed dose‐dependent binding of wild‐type R17 with an EC₅₀ of 160 nM, and negligible binding of the mutant R17^GAG2 (EC₅₀ > 1 μM; Figure  2A and S4, Supporting Information). Binding to BxPC3 cells was observed to be saturated with ≈500 nM of R17. In control experiments where cell surface HS was pre‐digested using a cocktail of heparinase enzymes (HSase, Figure S5, Supporting Information) or R17 was co‐incubated with heparin (5 μg mL⁻¹), we observed a significant reduction of R17 binding to cells (Figure 2B). These observations suggest that R17 binds HS GAGs and such binding events contribute largely in binding to BxPC3 cell surfaces.

Figure 2.

R17 binds to cell surface GAGs in Bxpc3s and causes the loss of HS GAGs. A) R17^WT and R17^GAG2 binding to fixed BxPC3 cells measured via flow cytometry. Data are shown as mean ± SD and represent two technical replicates. B) Flow cytometry measures R17^WT binding to fixed BxPC3 cells, which is abrogated with pre‐treatment with HS‐digesting heparinase enzymes (HSase) and in the presence of heparin (5 μg mL⁻¹). C) Representative histograms of cell surface HS levels as determined by flow cytometry after 24 hr of treatment with R17^WT. D) Calculated levels of normalized cell surface HS from C and other replicate data. Data are shown as mean ± SD for two independent experiments. E) Representative western blot of total HS levels in BxPC3 cells after 24 h treatment with R17^WT. F) Densitometry of normalized HS levels from e and other replicate data. Data are shown as mean ± SD for two independent experiments.

We next sought to evaluate the effects of R17 toward HS levels in live BxPC3 cells. We first confirmed that there were limited changes in the viability of these cells upon treatment with R17, in contrast to surfen, which displayed an IC₅₀ of 4 μM (Figure S6, Supporting Information). Using flow cytometry experiments of cells stained with an anti‐HS antibody clone 10E4,^{[ 26} , ^{27 ]} we observed a dose‐dependent loss of HS staining following the incubation of cells with R17 for 24 h, with a DC₅₀ of 467 nM (Figure 2C,D and S7, Supporting Information), which is similar to the concentration at which we observed binding saturation of R17 in BxPC3 cells (Figure 2A). We observed maximum degradation with a D _max of ≈60%, which was achieved at ≈2 μM of R17.

While flow cytometry experiments provided information on cell surface HS, we assessed the effects of R17 on total HS levels with western blotting. We observed that treatment of cells with R17 also caused the loss of total HS levels after 24 h (Figure 2E,F). These degradation effects were consistent with those observed when R17 was transiently overexpressed in BxPC3 instead of being added exogenously (Figure S8, Supporting Information). These results suggest that R17 binds cell surface HS and causes their loss from the cell surface and their eventual degradation.

HS chains are known to be degraded within the acidic compartments of lysosomes.^{[ 28 ]} Hence, we postulated that R17 could be causing the endocytosis of heparan sulfate proteoglycans (HSPGs) into lysosomes for eventual degradation, similar to other HS ligands.^{[ 20} , ²¹ , ^{22 ]} To demonstrate this, we performed confocal microscopy experiments with live cells over time to assess the localization of R17 to lysosomes. We observed that after 24 h of incubation, R17 does indeed significantly co‐localize with the lysosomal marker, Lysotracker (Figure  3A and Figure S9, Supporting Information). Using a logistic growth model, we determined that the half‐life of co‐localization was ≈6 h (Figure 3B). We also observed that R17 persists in cells in lysosomal compartments even after cells are washed (Figure S10, Supporting Information). To evaluate the dependence of R17 degradation on functional lysosomes, we co‐incubated cells with chloroquine, a weak base that accumulates within acidic lysosomal compartments and raises their pH.^{[ 29 ]} Addition of chloroquine to cells led to a modest increase in total HS levels and led to the loss of R17 efficacy to degrade HS (Figure 3C and S11, Supporting Information). We further investigated the lysosomal‐targeting abilities of R17 by evaluating the dependence of R17‐mediated HS degradation on one of the most well‐characterized lysosome‐targeting receptors‐ the cation‐independent mannose 6‐phosphate receptor (CI‐M6PR).^{[ 30} , ³¹ , ^{32 ]} Western blot analysis of total HS levels showed that R17 reduced HS levels even in the absence of CI‐M6PR, indicating that CI‐M6PR is not required for this mechanism (Figure 3D and S12, Supporting Information).

Figure 3.

R17 traffics to lysosomes and destines HS for lysosomal degradation. A) Confocal fluorescence microscopy images of live BxPC3 cells after 24 h treatment with R17. B) Co‐localization ratio of R17^{W T} with lysosomes quantified over 24 h time period using confocal microscopy, as shown in (A). C) Western blot analysis of total HS levels in BxPC3 cells after 24 h treatment with R17 in the presence or absence of chloroquine. D) Western blot analysis shows loss of total HS levels after 24 h treatment with R17 occurs at similar levels in WT and CI‐M6PR^KD BxPC3 cells.

To assess whether R17 modulates pancreatic cancer cell motility, we performed a scratch (wound‐healing) assay in BxPC3 cells. Relative to vehicle, R17 reduced wound closure over 24 h, indicating impaired lateral migration (Figure  4A). Quantification of wound width at matched fields demonstrated a dose‐dependent decrease in the reduction of closure (Figure 4B). To distinguish migration effects from potential contributions of cellular proliferation, a scratch (wound‐healing) assay was repeated with inclusion of Mitomycin C, a DNA‐crosslinking agent used to suppress proliferation (Figure 4C). Under these conditions, R17‐mediated decrease in wound closure persisted (Figure 4D), indicating the observed reduced closure is not due to altered proliferation. These data suggest that R17 attenuates cell migration in BxPC3 cells, consistent with a role for HS–dependent interactions in regulating cellular motility.^{[ 33} , ^{34 ]}

Figure 4.

R17 reduces BxPC3 cell migration. A scratch wound healing experiment was used to observe cell migration. A) Confluent layers of BxPC3 cells were manually scratched (outline indicates wound area) and observed over 48 h. Images of experiment showed increasing R17 concentrations inhibiting wound closure. B) Quantification of area in (A) contained within outline for 48 h timepoint. C) Confluent layers of BxPC3 cells were first treated with 5 μg mL⁻¹ of Mitomycin C for 2 h at 37 °C. Treated wells were scratched and subsequently treated with R17 with the indicated concentrations. Scratch closure was observed over 48 h. D) Quantification of area in (C) contained within outline for 48 h timepoint.

3. Conclusions

The endocytosis and trafficking of proteoglycans are well‐known to regulate the functions of cells, but how different GAG‐binding ligands influence such processes remains unclear.^{[ 4 ]} R17 is a protein among a panel of GAG‐binding ligands, and here we provide further evidence of its HS binding capabilities that appear to be similar to those reported for the Hedgehog protein.^{[ 35 ]} We demonstrate that R17 binding to HS on the surfaces of BxPC3 pancreatic cancer cells leads to its degradation. We observe that degradation is likely due to the trafficking of R17 to lysosomes in a manner that is independent of the CI‐M6PR. Intriguingly, R17 causes a reduction in the ability of BxPC3 cells to migrate, potentially due to its sequestration and clearance of chemokines. Since HS is known to play a significant role in tumor metastasis,^{[ 36} , ³⁷ , ^{38 ]} our data suggest that R17 reshapes the glycocalyx and likely alters the molecular landscape available for HS‐dependent interactions. While we have established the ability of R17 to bind HS in vitro and in cells, it remains to be understood whether R17 can have potent binding capabilities to other classes of GAGs (e.g., chondroitin sulfate, dermatan sulfate, or keratan sulfate).^{[ 39 ]}

Overall, our work highlights the need to continuously explore the effects of GAG‐binding ligands. Our work also asserts the emerging potential of using ligands for the induced endocytosis and lysosomal trafficking of HS and perhaps its associated cargo to regulate cellular functions.^{[ 13} , ²⁰ , ²² , ^{23 ]} As the field of extracellular protein degradation with lysosome‐targeting bifunctional molecules is growing,^{[ 32} , ⁴⁰ , ⁴¹ , ^{42 ]} the search for ligandable, lysosome‐directing cell surface receptors remains crucial for success.^{[ 43} , ^{44 ]} Given the ubiquity of HS, this pathway to traffic targets for lysosomal degradation may be an attractive alternative to classical means via the CI‐M6PR.^{[ 31 ]}

4. Experimental Section

4.1.

4.1.1.

Cell Culture

BxPC3 cells were purchased from ATCC and cultured in RMPI‐1640 (Gibco) supplemented with 10% v/v fetal bovine serum (FBS), 100 U mL⁻¹ penicillin, and 100 ug mL⁻¹ streptomycin. BxPC3 cells were grown at 37 °C in a humidified, 5% CO₂ incubator. Expi293F cells (Gibco) were grown in shaking incubators rotating at 135 rpm at 37 °C, 8% CO₂. Expi293F cells were cultured in Expi293 Expression Medium (Gibco) supplemented with 100 U mL⁻¹ penicillin and 100 ug mL⁻¹ streptomycin.

Expression of R17 and Constructs

Plasmid encoding R17^WT and R17^GAG2 (UniProt: E9M5R0 – AA: 28–439) was constructed as previously described (PMID: 24173234). Briefly, cDNA for R17^WT and R17^GAG2 were subcloned into the pHLsec (Addgene) vector with a N‐Terminal CD33 secretion signal C‐terminal 8xHis‐Tag. Both constructs were transiently expressed in Expi293F cells using ExpiFectamine 293 Transfection Kit (Gibco) according to the manufacturer's protocol. The culture medium was collected 8 days after transfection. Protein constructs were purified with cobalt resin (Thermo Scientific). Bound protein was washed with 30 mM imidazole and eluted with 300 mM imidazole. Eluted protein was then transferred into a dialysis membrane (Thermo Scientific) to allow for buffer exchange to 1× phosphate‐buffered saline (PBS) (overnight, 4 °C). From 30 mL of transfected Expi293F cells, we purified 0.9 mg of either R17^WT or R17^GAG2. Protein purity was confirmed via SDS‐PAGE shown in S9, Supporting Information.

Expression of Heparinase I, II, and III

Plasmids expressing heparinase I, II, or III from Flavobacterium heparinum (a gift from Prof. Jeffrey Esko) were transformed into BL21 (DE3) pLysS chemically competent E. coli cells. Selection was performed with ampicillin (100 μg mL⁻¹) agar plates for Heparinase II and III. Selection was performed with ampicillin (50 μg mL⁻¹), kanamycin (15 μg mL⁻¹), tetracycline (12.5 μg mL⁻¹), and chloramphenicol (200 μg mL⁻¹) agar plates for HSase I. Colonies were selected and grown overnight (37 °C, shaking) in starter cultures (5 mL lysogeny broth (LB) with corresponding antibiotic(s)). For each construct, LB (1 L) with the corresponding antibiotic was inoculated with starter culture (3 mL) and grown to OD₆₀₀ ≈0.6. Each culture was induced (1 mM IPTG, O/N, 37 °C, shaking) and harvested (3,600 ×g, 15 min, 4 °C) separately. The resulting cell pellets were lysed with a Branson sonicator (1 s on, 3 s off, 3 min total, 20% amplitude) in wash buffer (25 mM imidazole, 3× PBS,pH 7.4) containing protease inhibitor cocktail and pre‐clarified (16 000 ×g, 30 min, 4 °C). The supernatants were filtered with 0.45 μm filter, and 2 mL of pre‐washed Cobalt beads were added (O/N, 4 °C, rocking). Lysates were loaded onto separate gravity columns and washed five times (20 mL, 25 mM imidazole, 3× PBS, pH 7.4) then eluted (250 mM imidazole, 3× PBS, pH 7.4). Isolated proteins were desalted via PD10 column and concentrated in 1× PBS using centrifugal filters (Amicon Ultra Centrifugal Filter, 10 kDa MWCO). Protein concentration was determined by the DC Protein Assay (Bio‐rad), and purity was confirmed by a stain‐free gel shown in S10, Supporting Information. Equimolar protein was combined, and the activity was validated on free heparin sulfate by measuring A₂₃₅ via NanoDrop One^C.

Confocal Imaging and Quantitative Analysis

BxPC3 cells were seeded in 35‐mm glass‐bottom dishes (MatTek Corporation) and allowed to adhere overnight. The next morning after seeding, lysosomes were labeled using Lysotracker Deep Red (Invitrogen, 1:20 000 dilution in culture medium, 30 min, 37 °C). After a 30 min incubation, cells were washed with culture medium three times. Hoescht dye was added to cells (1:1000 in culture medium, 10 min, RT) and washed 3× to visualize cell nuclei. Then, 100 nM R17 was pre‐complexed in culture medium with mouse anti‐6x His (Alexa Fluor 555, Invitrogen, cat. MA1‐21315‐A555) in a 1:1 molar ratio for 30 min before being added to cells.

Two‐color confocal imaging was performed using a Zeiss LSM 880 laser scanning confocal microscope (Carl Zeiss, Oberkochen, Germany) equipped with a Plan‐Apochromat 40X oil‐immersion objective (NA 1.4). Red fluorescent protein (RFP); (corresponding to R17: anti‐6x His Alexa Fluor 555 Ab) was excited using the 561 nm line of a diode‐pumped solid‐state laser, and emission was collected through a 580–630 nm band‐pass filter. Cy5 (Lysotracker Deep Red labeled lysosomes) was excited using the 647 nm line of a helium‐neon (HeNe) laser, and emission was collected through a 655–680 nm band‐pass filter. To avoid spectral bleed‐through, channels were acquired sequentially with optimized laser power and detector gain for each fluorophore. All images were recorded using identical acquisition parameters, including a pixel size of 0.07 μm pixel⁻¹, ensuring precise spatial registration between channels.

For quantitative image analysis, bright fluorescent puncta in each fluorescence channel were detected and counted on a per‐cell basis using the Find Maxima function in Fiji (version 2.14.0), followed by conversion to binary masks. The total signal area from the RFP channel ($S_{\mathrm{R}17}$) was defined as the number of pixels containing the R17 signal. Similar analysis for the Cy5 channel was used to quantify total signal area for lysosomes ($S_{\text{Lysosome}}$). To quantify co‐localization with lysosomes, binary images from RFP and Cy5 channels were multiplied using the Image Calculator function in Fiji, and the total overlapping pixel area ($I(t)$) was measured. The fraction of R17 signal colocalized with lysosomes was expressed as the ratio to total R17 area

$$\text{Ratio}\hspace{0.17em}\text{of}\hspace{0.17em}\text{total}\hspace{0.17em}\mathrm{R}17:I(t)=\frac{\sum (S_{\mathrm{R}17}\times S_{\text{Lysosome}})}{\sum (S_{\mathrm{R}17})}$$ (1)

This quantification was performed for each time point (6, 7, 8.5, and 24 h).

To model the time‐dependent increase in R17‐lysosome overlap, we applied a logistic growth model, a framework commonly used to describe systems exhibiting self‐limited growth due to resource constraints.^{[ 45} , ^{46 ]} In this context, the overlap ratio initially increases rapidly as available R17 and lysosomes interact, but the rate slows as the number of unbound components decreases, ultimately reaching a steady‐state plateau. The temporal evolution of the overlapping fraction($I(t)$) was fitted using the logistic equation

$$I(t)=\frac{I_{\text{Max}}}{1+{\mathrm{e}}^{-k\left(t-t_{\frac{1}{2}}\right)}}$$ (2)

where $I_{\text{Max}}$ represents the maximum overlap ratio, $k$ is the intrinsic overlap rate constant, and $t_{\frac{1}{2}}$ is the time to reach half of the maximum overlap.

Cytotoxicity Assay

Cells were seeded in 96‐well plates at a density of 0.01×10⁶ cells mL⁻¹ and allowed to adhere overnight at 37 °C, 5% CO₂. Medium was replaced with treatment medium containing test compounds (R17, surfen, etc.) or vehicle controls across a serial dilution series and incubated for the respective times. After appropriate incubation, plates were equilibrated to room temperature (RT) for 20 min, and cell viability was quantified using the Promega CellTiter‐Glo Luminescent Viability Assay (Promega). CellTiter‐Glo reagent was added at an equal volume to the amount of cell culture medium present in each well (e.g. 100 μl reagent added to 100 μl of medium). Contents were mixed for 2 min on an orbital shaker and incubated at RT for 10 min to stabilize the luminescence signal. Luminescence was recorded on a plate reader with no filter. Raw signals were blank‐subtracted (medium‐only wells) and normalized to intra‐plate controls: percent viability = (sample/vehicle control) × 100. Dose–response curves (viability versus concentration) were fit to a logistic model in GraphPad Prism to derive IC₅₀/EC₅₀ values. All treatments were performed in technical triplicate unless otherwise indicated.

Scratch (Wound‐Healing) Assay

The scratch migration assay was performed in 12‐well plates. Briefly, BxPC3 cells (initially cultured in roswell park memorial institute (RPMI)‐1640 supplemented with 10% v/v FBS, 100 U mL⁻¹ penicillin, 100 ug mL⁻¹ streptomycin) were detached using an enzymatic dissociation reagent, pelleted (500 × g, 3 min), washed once with PBS, and resuspended in fresh growth medium. Cells were seeded into 12‐well plates at a density predicted to reach ≈80% confluence after 24 h, then changed to serum‐free medium (1–2 mL per well) and cultured for an additional 24 h. In the scratch assay with mitomycin C, cells were treated with 5 μg mL⁻¹ Mitomycin C proliferation inhibitor (10 mg mL⁻¹ stock, Sigma) for 2 h at 37 °C, 5% CO₂. Wells were washed with 1× PBS three times to remove excess Mitomycin C. To create the wound, the confluent monolayer was scratched across the well diameter with a 200 μL pipette tip in contact with the plastic, taking care not to gouge the surface. Wells were gently washed with PBS to remove detached cells and debris, replenished with serum‐free medium (1–2 mL per well) with or R17, and immediately imaged on a light microscope equipped with a digital camera (EVOS M5000, Invitrogen). R17 was titrated at 2000, 500, and 0 nM. For each well, fields spanning intact monolayer on both sides of the wound were recorded, and the plate underside was marked to enable re‐imaging of the same positions. Plates were returned to the incubator for 24 h, after which the same fields were re‐imaged; for longer time courses, medium/treatments were refreshed as needed and additional images acquired at the indicated times. Wound width was quantified in ImageJ/Fiji from the immediate post‐scratch image (a) and the matched later image (b), and percent closure was calculated as ((a−b)/a) × 100.

ELISA: Binding ELISA

Assays were performed in heparin‐coated 96‐well microplates (Bioworld). Unless otherwise indicated, incubations were at RT, and volumes refer to per‐well amounts. Phosphate‐buffered saline with 0.2% v/v Tween‐20 (PBST) was used for all washes, and PBST supplemented with 1%–2% w/v bovine serum albumin (BSA) served as diluent and blocking buffer. Wells were blocked with 200 μL PBST + 2% BSA for 1 hr, then washed 5× with 200 μL PBST per wash. R17 (His‐tagged) was added in PBST + 1% BSA (50 μL per well) as a titration series (500, 250, 125, 62.5, 31.25, 15.625, and 0 nM) and incubated for 1 h. Plates were then washed 5× with PBST (200 μL per wash) and incubated with primary antibody diluted in PBST + 1% BSA (50 μL per well, 1 h); mouse anti‐His was used at 1:1000 (BioLegend, cat. 652 501). Following 5× PBST washes (200 μL per wash), wells were incubated with horseradish peroxidase (HRP)‐conjugated goat anti‐mouse IgG secondary antibody (Invitrogen, cat. G21040) at 1:1000 in PBST + 1% BSA (50 μL per well, 1 h), then washed again 5× with PBST (200 μL per wash). ELISA substrate (BioFX TMB One Component HRP Microwell Substrate) was equilibrated to RT and added at 50 μL per well; color was developed for 5–20 min at RT, and the reaction was quenched with 1 m sulfuric acid (100 μL per well). Absorbance was measured at 450 nm on a microplate reader.

ELISA: Competition ELISA

Assays were performed in heparin‐coated 96‐well microplates (Bioworld). Unless otherwise indicated, incubations were at RT, and volumes refer to per‐well amounts. Phosphate‐buffered saline with 0.2% v/v Tween‐20 (PBST) was used for all washes, and PBST supplemented with 1%–2% w/v BSA served as diluent and blocking buffer. Wells were blocked with 200 μL PBST + 2% BSA for 1 hr, then washed 5 × with 200 μL PBST per wash. The first ligand—either R17 at 33 nM final concentration or surfen at 9 μM—was added in PBST + 1% BSA (50 μL) and incubated 10–15 min. Without washing, the second ligand (surfen or R17, respectively) was titrated and co‐incubated with the first ligand for 1 h. R17 was titrated at 500, 250, 125, 62.5, 31.25, 15.625, and 0 nM; surfen was titrated at 20, 10, 5, 2.5, 1.25, and 0 μM. Plates were then washed 5× with PBST (200 μL per wash) and incubated with primary antibody diluted in PBST + 1% BSA (50 μL per well, 1 h); mouse anti‐His was used at 1:1000 (BioLegend, cat. 652 501). Following 5× PBST washes (200 μL per wash), wells were incubated with HRP‐conjugated goat antimouse IgG secondary antibody (Invitrogen, cat. G21040) at 1:1000 in PBST + 1% BSA (50 μL per well, 1 h), then washed again 5× with PBST (200 μL per wash). ELISA substrate (BioFX TMB One Component HRP Microwell Substrate) was equilibrated to RT and added at 50 μL per well; color was developed for 5–20 min at RT, and the reaction was quenched with 1 m sulfuric acid (100 μL per well). Absorbance was measured at 450 nm on a microplate reader.

Flow Cytometry Experiments: R17 ^WT and R17 ^GAG2 Binding to Fixed BxPC3 Cells Detected by Flow Cytometry

BxPC3 cells were seeded in 12‐well plates in culture medium and grown to 80% confluency over 36 h. For conditions using BxPC3 cells pre‐digested with heparinases, an equimolar mixture of heparinases I, II, and III was added to cells 16 h after seeding, diluted at 1:10 000 in culture medium (total 1 mL). Cells were incubated with the heparinase mixture for 24 h. The BxPC3 cells were then dissociated using 200 μL Accutase (Biolegend) and rinsed three times with 200 μL 1% BSA in RPMI medium (buffer). All subsequent washes and incubations were done with a total buffer volume of 200 μL. Cells were incubated with 3% paraformaldehyde (PFA) in PBS (15 min, rt). After two washes with buffer, the R17 protein construct was added to the fixed cells in buffer (1 h, 4 °C). For the competition assay with heparin, heparin at a concentration of 5 ug mL⁻¹ (Galen Molecular, cat. HEP001) was pre‐mixed with a solution of R17 and then added to fixed cells in buffer (1 h, 4 °C). The cells were then rinsed with buffer twice to remove unbound R17, followed by incubation with anti‐6X‐His antibody (Invitrogen, cat. MA1‐21 315‐A555) (1:500) (1 h, 4 °C). After washing the stained cells twice with buffer, data were collected on NovoCyte Quanteon 4025 with NovoSampler using NovoExpress software. Each cell population analyzed for median fluorescence intensity (MFI) contains at least 10 000 gated events. Sample gating is shown in S11, Supporting Information.

Flow Cytometry Experiments: R17‐Mediated HS Degradation of BxPC3 Cells Detected by Flow Cytometry

BxPC3 cells were seeded in 12‐well plates in culture medium and allowed to adhere overnight. The following morning, roughly 16 h after seeding, the cells were dosed with R17 in culture medium (total 1 mL). After 24 h of incubation with R17, BxPC3 cells were dissociated using 200 μL Accutase (Biolegend) and then washed three times with 1% BSA in RPMI medium (buffer). All subsequent washes and incubations were done with a total buffer volume of 200 μL. Cells were incubated with anti‐HS antibody (Amsbio, clone: 10E4) (1:200) (1 h, 4 °C). Cells were washed with buffer two times. The bound 10E4 was labeled with goat anti‐mouse IgM (Invitrogen, cat. A21238) (1:500) in buffer (1 h, 4 °C). After two rinses with buffer, cells were fixed with 3% PFA in PBS (15 min, rt). Data was collected on either of the following two instruments: NovoCyte 3000 with NovoSampler Pro using software BD FACS Diva 6 or NovoCyte Quanteon 4025 with NovoSampler using NovoExpress software. Data were analyzed using FlowJo v.10.10.0 for Mac. Each cell population analyzed for MFI contains at least 10 000 gated events. Sample gating is shown in S12.

SiRNA‐Mediated Knockdown

BxPC3 cells were reverse‐transfected at the time of seeding using Lipofectamine RNAiMAX (Thermo Fisher Scientific, cat. 13 778 075) according to manufacturer's protocol. Cells were seeded in 6‐well plates in culture medium with 30 pmol of CI‐M6PR‐directed siRNA and 18 uL of Lipofectamine RNAiMAX (pre‐incubated 5 min, RT in Opti‐MEM). Cells were seeded to achieve 80% confluency post‐seeding, as transfected cells were treated with R17 for 24 h.

Western Blotting

BxPC3 cells were seeded in 6‐well plates in culture medium. After 16 h, cells were dosed with R17 in culture medium. For experiments using endocytosis or lysosomal inhibitors, compounds were dosed in media containing 0.1% DMSO. Cell lysates were boiled in 1× Laemmli Buffer (Bio‐rad) with 10% β‐mercaptoethanol and loaded on to 4%–15% TGX‐PAGE gels (Bio‐rad). Gels were then transferred to PVDF membranes using the Trans‐Blot Turbo Transfer system (Bio‐rad) according to manufacturer protocols. The membranes were blocked with 5% BSA in TBST (1 h, RT), then incubated with primary antibody in 5% BSA in TBST (overnight, 4 °C) (GAPDH antibody (Biolegend, 1:5000); HS antibody (Amsbio, clone:10E4) (1:100); CI‐M6PR antibody (Cell Signaling, clone: D3V8C) (1:1000)). Blots were washed three times with TBST followed by incubation with secondary antibody in 5% BSA in TBST (1 h, RT) (Goat anti‐rat IgG (Biolegend, clone: Poly4054) (1:10 000); Goat anti‐Mouse IgM (Sigma Aldrich, cat. AP500P) (1:5000); and Goat anti‐Rabbit IgG (Invitrogen, cat. A32733)).

Supporting Information

Supporting Information is available from the Wiley Online Library or from the author.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Supplementary Material

Data Availability Statement

The data that support the findings of this study are available in the supplementary material of this article.