PKC-Delta-Dependent Pathways Contribute to the Exacerbation of the Platelet Activity in Crohn's Disease

The pathogenesis of inflammatory bowel diseases (IBD), representing chronic and relapsing-remitting disorders of the gastrointestinal tract, encompasses Crohn's disease (CD) and ulcerative colitis.[1] [2] Although the cause and mechanisms of both conditions remain unknown, mounting evidence suggests that gut tissue injury is not exclusively the result of a dysregulated immune response, but also involves the active participation of multiple non-immune cellular systems.[3] [4]

In addition to the well-described classical function of platelets in primary hemostasis, there are a rising number of studies supporting their significant role as amplifying agents in inflammatory processes/disorders and immune response,[5] [6] [7] through the release a broad spectrum of pro-inflammatory mediators.[8] [9] [10] [11] [12]

Platelet activation by multiple signaling pathways induces their shape change and release of intraplatelet stored granules. Members of the protein kinase C (PKC) family, highly expressed in platelets, are important mediators of these processes.[13] [14] In particular, conventional and novel PKC isoforms were previously identified as a central and redundant element of platelet signaling pathways with distinct functions in platelet granule release (PKCα, ε, δ, θ).[13] [15] [16]

PKCδ plays a pivotal role in growth regulation and tissue remodeling in other cells[17] and differentially regulates platelet function, depending on the platelet agonists and their receptors.[18] [19] [20] It has been shown that PKCδ^−/− mice have more platelets and megakaryocytes than wild-type littermate control mice.[21] However, the underlying mechanism of hyperactivated platelets and the contribution of platelet-PKCδ in the mechanisms of inflammation, in CD, are not completely understood.

In this study, pharmacological and molecular genetic approaches were used to investigate the functional role of PKCδ isoform and downstream effectors in modulation of molecular inflammatory mechanisms during CD pathogenesis.

In human platelets, pre-treatment with the specific PKCδ inhibitor significantly decreased platelet activation in patients with CD. Analysis of PKCδ phosphorylation indicates that it is positively regulated by the mitogen-activated protein kinase (MAPK) pathway.

These findings not only constitute a potential to explain the mechanism of hyperactivated platelets in patients with active CD, but also confirm that PKCδ is an important signal transducer of multiple signaling pathways,[22] [23] showing for the first time PKCδ as an appropriate therapeutic target for the treatment of chronic intestinal inflammation pathogenesis.

A total of 89 of CD patients were recruited from CHU Ibn Rochd Casablanca between October 15, 2018 and December 31, 2019. The diagnosis of CD was made based on a combined assessment of abdominal imaging, endoscopy, symptomatology, and histology. The median (interquartile range) age of CD patients (47 [62–27] years) was matched with healthy donors (42 [65–33] years). All patients and healthy donors (HD) gave written informed consent. Furthermore, all experiments using human subjects were performed in accordance with the Declaration of Helsinki. The study was evaluated and approved by the Ethics Committee of the Faculty of Medicine, Casablanca-Hassan II University (No. PR-0304–18).

Antibodies against phospho-PKCδ (Tyr³¹¹ and Tyr⁵⁰⁵) and total PKCδ were purchased from Cell Signaling Technology (Beverly, MA). Bovine thrombin was purchased from Sigma-Aldrich (Oaskville, ON), while native type I collagen was from Chronolog Corp. The p38 SB203580 inhibitor was obtained from Calbiochem (San Diego, CA).

PKCδ membrane activity/translocation was selectively inhibited by the peptide antagonist δ(V1–1)TAT as previously described.[24] Briefly, the PKCδ δ(V1–1) interacting sequence of the receptors for activated C kinases (SFNSYELGSL) was coupled to the TAT HIV membrane permeable sequence (YGRKKRRQRRR: amino acids 47–57 of TAT) through a cysteine–cysteine bridge. Peptide sequencing was performed at CanPeptide Inc. (Pointe-Claire, QC).

Cytokine levels in plasma and platelets were assessed using Bio-Plex Pro Human Cytokine 27-plex Assay (Bio-Rad, Marnes-la-Coquette, France). If cytokines were not detected, a concentration corresponding to the lowest value extrapolated from the standard curve was attributed to the sample for quantification and comparison analyses.

To prepare human platelets, fresh venous blood was collected from healthy volunteers, free from medications known to interfere with platelet function, at least 10 days before the experiment. Washed platelets were prepared as previously described.[25] [26] Briefly, platelet-rich plasma (PRP) was obtained by centrifugation of acid citrate dextrose (ratio of 1:5) anticoagulated blood at 200 g for 15 minutes. Platelets were then pelleted from PRP, to which 1 μg/mL of PGE₁ was added, washed with HBSS-Hank's sodium citrate buffer (138 mM NaCl, 5 mM KCl, 0.34 mM Na₂HPO₄, 0.4 mM KH₂PO₄, 4.2 mM Na₂HCO₃, 5.6 mM Glucose, 10 mM HEPES, 12.9 mM sodium citrate, pH 7.4), also containing PGE₁ (0.5 μg/mL), and finally resuspended in HBSS-Hank's buffer containing 2 mM MgCl₂ and 2 mM CaCl₂. Platelets were adjusted to 250 × 10⁶ /mL and allowed to rest at 37°C for 30 minutes before further manipulation.

Platelet aggregation was then monitored on a four-channel optical aggregometer (Chrono-log Corp.) under shear conditions (1,000 rpm) at 37°C in the presence of thrombin or collagen. Traces were recorded until stabilization of platelet aggregation was reached.

Platelets were stimulated with thrombin in the absence or presence of δ(V1–1)TAT or SB203580 for the appropriate time under shear conditions at 37°C. The reaction was stopped by adding the appropriate volume of 4 x SDS-Laemmli buffer. Platelet lysates were heated for 5 minutes at 95°C and analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE).

Proteins were resolved in 8% SDS-PAGE gels and transferred onto nitrocellulose membranes. The membranes were blocked with 5% nonfat dry milk for 1 hour, washed three times with TBS/Tween (150 mM NaCl, 20 mM Tris, pH 7.4, 0.1% Tween-20), and incubated overnight at 4°C with the indicated antibody. Following washing steps, membranes were labeled with horseradish peroxidase-conjugated secondary antibody for 1 hour, washed and bound peroxidase activity was then detected by enhanced chemiluminescence (PerkinElmer Life Sciences, Waltham, MA). To assess equal amounts of protein loading, membranes were stripped, blocked with 5% milk, and blotted for total PKCδ.

For confocal immunofluorescence and imaging, freshly isolated platelets were fixed with paraformaldehyde 2% (v/v) and allowed to immobilize on poly-L-lysine-coated coverslips. Adhered platelets were permeabilized for 20 minutes with Triton X-100 containing 2% bovine serum albumin (BSA). Platelets were then incubated for 3 hours with the rabbit anti-human polyclonal anti-PKCδ antibody (Abcam) and the mouse anti-human monoclonal antibody against α-tubulin (Santa Cruz Biotechnology) to delineate the membrane, washed and labeled with anti-rabbit IgG–Alexa 555 and anti-mouse IgG Alexa-488 secondary antibodies for 1 hour at room temperature. Coverslips were mounted on microscopic slides and a series of fluorescent confocal images (Z-stacks) were acquired using a LSM 510 confocal microscope (Zeiss, Oberkochen, Germany).

Scanning electron microscopy platelets from healthy subjects and CD patients stimulated with thrombin (0.02 and 0.5 U/mL) were allowed to immobilize on 2% BSA-treated glass coverslips for 30 minutes at 37°C. Samples were then fixed in 2% paraformaldehyde overnight at 4°C. Dehydration of surfaces was achieved by placing samples in ethanol/water followed by amyl acetate/ethanol baths for 15 minutes each, increasing the ethanol/water proportion from 30 to 100% and the amyl acetate/ethanol proportion from 25 to 100%. Slides were subsequently coated with gold palladium particles and analyzed on a Hitachi S-4700 Field Emission Gun Scanning Electron Microscope. For granule secretion, P-selectin (CD62P) translocation to the platelet surface from α-granule stores was measured by flow cytometry. Briefly, human platelets were stimulated with thrombin, fixed, washed, and stained with saturating concentrations of anti-CD62P PE-conjugated antibody (BD Biosciences). Platelets were analyzed on an Altra flow cytometer (Beckman Coulter).

The level of serum amyloid A (SAA) in plasma was measured by sandwich ELISA using a commercial kit (E-90SAA, Immunology Consultants Laboratory, Inc., Portland). SAA ELISAs were performed following the manufacturer's instructions.

Statistical comparisons were done using a one-way ANOVA, followed by a Dunnett's-t-test for comparison against a single group. The values with p ≤0.05 were considered statistically significant. The data and statistical analysis comply with the recommendations on experimental design and analysis in pharmacology.[27]

Using a multiplex assay to monitor 27 cytokines in plasma prepared from healthy volunteers (n = 10) and from patients with CD (n = 44), we determined that the concentrations of eight cytokines or growth-factors were significantly increased (granulocyte colony-stimulating factor [G-CSF], interferon gamma [IFN-γ], IL-1β, interferon gamma-induced protein 10 [IP-10], macrophage inflammatory proteins 1β [MIP-1β], macrophage-derived chemokine [MDC], regulated on activation, normal T cell expressed and secreted [RANTES], TNFα [transforming growth factor β], vascular endothelial growth factor A [VEGF-A]); the concentrations of two cytokines (IL-9 and IL-15) were significantly decreased, while that of 17 cytokines or growth-factors were similar to healthy controls ([Fig. 1A]). Elevated inflammatory cytokine levels are reported in the blood of patients with CD and may contribute to the overwhelming inflammation.

Fig. 1 Platelet hyper-reactivity in patients with CD is PKCδ-dependent. (A) Heat map visualization of 27 cytokine expression profiles in plasma with CD (median of n = 44) versus healthy donors (median of n = 10). Cytokine expression is represented as a (log2) fold change relative to healthy donors. The numbers in each part represent the change and statistical significance (in brackets), and the color codes refer to blue for increased expression and pink for decreased expression. (B) Heat map visualization of 23 cytokine expression profiles in plasma of patients with CD (median of n = 44) relatively to the heathy donors (median of n = 10). Cytokine expression is represented as a (log2) fold change relative to healthy donors. The numbers in each part represent the change and statistical significance (in brackets) and the color codes refer to blue for increased expression and pink for decreased expression. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. (C) Low doses of thrombin induce platelet shape change. Scanning electron micrographs of α-thrombin-stimulated platelets (0.02 U/mL) at low (top: ×2500) and high (bottom: ×30,000) magnifications. Images are representative of three independent experiments. (D) Platelets from CD patients are more reactive at low doses of α-thrombin and collagen. Low doses of α-thrombin and collagen potentiate platelet aggregation in platelets from CD patients versus healthy donors. Histogram represents the mean of data ± SD of aggregation traces (n = 10, p < 0.001). (E) Representative traces of platelet aggregation induced by a priming dose (0.02 U/mL) of α-thrombin. Platelets were stimulated with 0.02 U/mL of α-thrombin (traces on the left) or 0.5 μg/mL of collagen (traces on the right). (F) PKCδ is translocated to the membrane in patients with CD, in response to a priming dose of α-thrombin (0.02 U/mL). Representative confocal microscopy images showing platelets double stained for PKCδ (red) and α-tubulin (green) taken at 63X magnification. (G) Phosphorylation of PKCδ on Tyr³¹¹ (but not on Thr⁵⁰⁵) is decreased in CD patients following PKCδ and p38 MAPK inhibition, in response to a priming dose of α-thrombin. Platelets were pre-treated with the specific PKCδ inhibitor (δ(V1–1)TAT, 1 μM) and/or p38 MAPK inhibitor (SB203580, 10 μM) for 5 minutes at 37°C under shear conditions. Aggregation was then initiated by a priming dose (0.02 U/mL) of α-thrombin and the reaction was terminated at 5 minutes. Platelet lysates were then analyzed by SDS-PAGE for p-PKCδ Thr⁵⁰⁵ and p-PKCδ Tyr³¹¹ (n = 5). Blots for total PKCδ are from a representative stripped membrane.

We also monitored cytokines in lysates prepared using platelets from patients with CD and included healthy volunteers as controls. While a few cytokines were below the detection limit (G-CSF, MIP-1α, MIP-1β, and VEGF-A), we detected 23 cytokines/growth-factors. Noteworthy, we found significantly reduced levels for six cytokines/growth-factors including cytokines relevant to inflammatory responses (e.g., IFNγ, IL-1β, and TNF α; [Fig. 1B]) in CD. These data show an increase in platelet-derived cytokines in plasma and a parallel decrease in cytokines in platelets themselves, suggesting more release from platelets in CD.

As shown in [Fig. 1C], platelets from patients with CD are more reactive than control platelets, in response to minimal doses of thrombin. This hyper-reactivity observed in platelets from CD patients, was displayed by a significant morphological shape change characterized by an increase in lamellipodia and filopodia formation and also by increased α-granule secretion ([Fig. 2A]).

Fig. 2 (A) α-granule secretion (percent of CD62P positive platelets) is increased in response to a priming dose of thrombin (0.02 U/mL) in CD patients versus healthy subjects. No significant difference was observed between these two groups in response to high doses of thrombin (0.5 U/mL). Histogram represents the mean of data ± SD of plots for CD62P expression (n = 3) ***p <0.05. (B) Phosphorylation of PKCδ on Tyr³¹¹ and Thr⁵⁰⁵ is not affected in healthy subjects following PKCδ and p38 MAPK inhibition, in response to a priming dose of α-thrombin. Platelets were pre-treated with the specific PKCδ inhibitor (δ(V1–1)TAT, 1 μM) and/or p38 MAPK inhibitor (SB203580, 10 μM) for 5 minutes at 37°C under shear conditions. Aggregation was then initiated by a priming dose (0.02 U/mL) of α-thrombin and the reaction was terminated at 5 minutes. Platelet lysates were then analyzed by SDS-PAGE for p-PKCδ Thr⁵⁰⁵ and p-PKCδ Tyr³¹¹ (n = 5). Blots for total PKCδ are from a representative stripped membrane.

Previous studies have identified the critical role of PKCδ in platelet activation.[18] [19] [20] [28] To confirm the hyper-reactivity of platelets from CD patients, the aggregation test was performed. As anticipated, low doses of α-thrombin (0.02 U/mL) and collagen potentiate platelet aggregation in platelets from CD patients versus healthy subject ([Fig. 1D] and [E]).

We then evaluated the translocation of PKC delta to the membrane. As shown in [Fig. 1F], PKCδ is translocated to the membrane in patients with CD, in response to a priming dose of α-thrombin.

To further explore the signaling mechanism involved in the regulatory role of PKCδ in thrombin-induced platelet aggregation in CD patients, we assessed the phosphorylation of PKCδ on Tyr³¹¹ and Thr⁵⁰⁵. Phosphorylation of PKCδ on Tyr³¹¹ (but not on Thr⁵⁰⁵) is decreased following PKCδ and p38 MAPK inhibition, in response to a priming dose of α-thrombin (0.01 U/mL) ([Fig. 1G]).

Although platelets are traditionally recognized for their contribution to the pathology of disorders related to thrombosis, many lines of research clearly demonstrate that these immune cells also contribute to inflammation.[8] [29] [30]

In the pilot study including 89 patients with CD (median age 47 years) and 34 age-matched HD, we have evaluated the role of PKCδ in platelet activation in CD. Our results demonstrate for the first time that PKCδ positively regulates, under inflammatory conditions, platelet function upstream of p38 MAPK pathway, in response to thrombin.

It has been previously shown that PKCδ isoform can negatively and positively regulate platelet activation, secretion, and aggregation, in response to different platelet agonists, including thrombin and collagen.[18] [31] [32] [33] Phosphorylation of the PKC family is an essential regulatory mechanism required for complete kinase activity. Particularly, PKCδ isoform is regulated through extensive multi-phosphorylations on threonine, serine, and tyrosine residues.[34] [35] [36] Initially, we recorded a translocation of PKCδ from the cell cytosol to different subcellular compartments in response to low doses of thrombin, in platelets from CD patients versus healthy subjects. This phenomenon, as we have demonstrated, is associated with a potentiation of aggregation and a change of form. Thus, we focused, in this brief report, on the evaluation of PKCδ phosphorylation at the two key residues (Tyr³¹¹ and Thr⁵⁰⁵) following downstream activation of G-protein-activated pathways in platelets. Our data show that phosphorylation of PKCδ on Tyr³¹¹ in CD patients (but not on Thr⁵⁰⁵) is significantly decreased, following specific inhibitions of PKCδ and p38 MAPK, in response to a priming dose of α-thrombin. In platelets from Healthy Subjects, PKCδ phosphorylation on Tyr³¹¹ and Thr⁵⁰⁵ appears to be unaffected by δ(V1–1)TAT and SB203580 treatments ([Fig. 2B]).

Recently, it has been shown that PKCδ protein expression is enhanced during megakaryocyte differentiation. Indeed, deletion of PKCδ in mice caused an increase in platelet production probably due to an enhanced megakaryocyte production in the bone marrow and spleen, suggesting that PKCδ isoform is a major intraplatelet regulator.[21]

In conclusion, these studies provide a clear indication that PKCδ Tyr³¹¹ phosphorylation is an important molecular regulator during inflammation. Our results indicate that the PKCδ/p38 MAPK axis can damper platelet activation in the presence of suboptimal thrombin concentrations. Overall, these findings indicate that regulating the PKCδ pathway may provide novel therapeutic strategies to treat non-resolving inflammatory pathologies in humans.

Authors' Contributions

Y.L., N.S., and Y.Z. designed and performed the major of experiments, analyzed data, and assisted with writing the manuscript. N.Z. and L.K. participated in the recruitment of patients. A.N., N.H., F.J., M.O., and Y.Z. designed experiments, analyzed data, and prepared the manuscript. All authors read and approved the final manuscript.

Publication History

Article published online:
08 November 2021

• References

• 1 Senhaji N, Nadifi S, Zaid Y. et al. Polymorphisms in oxidative pathway related genes and susceptibility to inflammatory bowel disease. World J Gastroenterol 2017; 23 (47) 8300-8307
• 2 Marafini I, Troncone E, Salvatori S, Monteleone G. TGF-β activity restoration and phosphodiesterase 4 inhibition as therapeutic options for inflammatory bowel diseases. Pharmacol Res 2020; 155: 104757
• 3 Fiocchi C. Inflammatory bowel disease: etiology and pathogenesis. Gastroenterology 1998; 115 (01) 182-205
• 4 Fiocchi C. Intestinal inflammation: a complex interplay of immune and nonimmune cell interactions. Am J Physiol 1997; 273 (04) G769-G775
• 5 Weyrich AS, Lindemann S, Zimmerman GA. The evolving role of platelets in inflammation. J Thromb Haemost 2003; 1 (09) 1897-1905
• 6 Semple JW, Italiano Jr JE, Freedman J. Platelets and the immune continuum. Nat Rev Immunol 2011; 11 (04) 264-274
• 7 Allaoui A, Khawaja AA, Badad O. et al. Platelet Function in Viral Immunity and SARS-CoV-2 Infection. Semin Thromb Hemost 2021; 47 (04) 419-426
• 8 Semple JW, Freedman J. Platelets and innate immunity. Cell Mol Life Sci 2010; 67 (04) 499-511
• 9 Jenne CN, Urrutia R, Kubes P. Platelets: bridging hemostasis, inflammation, and immunity. Int J Lab Hematol 2013; 35 (03) 254-261
• 10 Senhaji N, Kojok K, Darif Y, Fadainia C, Zaid Y. The Contribution of CD40/CD40L Axis in Inflammatory Bowel Disease: An Update. Front Immunol 2015; 6: 529
• 11 Zaid Y, Guessous F, Puhm F. et al. Platelet reactivity to thrombin differs between patients with COVID-19 and those with ARDS unrelated to COVID-19. Blood Adv 2021; 5 (03) 635-639
• 12 Zaid Y, Puhm F, Allaeys I. et al. Platelets Can Associate with SARS-Cov-2 RNA and Are Hyperactivated in COVID-19. Circ Res 2020
• 13 Harper MT, Poole AW. Diverse functions of protein kinase C isoforms in platelet activation and thrombus formation. J Thromb Haemost 2010; 8 (03) 454-462
• 14 Heemskerk JW, Harper MT, Cosemans JM, Poole AW. Unravelling the different functions of protein kinase C isoforms in platelets. FEBS Lett 2011; 585 (12) 1711-1716
• 15 Hall KJ, Harper MT, Gilio K, Cosemans JM, Heemskerk JW, Poole AW. Genetic analysis of the role of protein kinase Ctheta in platelet function and thrombus formation. PLoS One 2008; 3 (09) e3277
• 16 Pears CJ, Thornber K, Auger JM. et al. Differential roles of the PKC novel isoforms, PKCdelta and PKCepsilon, in mouse and human platelets. PLoS One 2008; 3 (11) e3793
• 17 Steinberg SF. Distinctive activation mechanisms and functions for protein kinase Cdelta. Biochem J 2004; 384 (Pt 3): 449-459
• 18 Yacoub D, Théorêt JF, Villeneuve L. et al. Essential role of protein kinase C delta in platelet signaling, alpha IIb beta 3 activation, and thromboxane A2 release. J Biol Chem 2006; 281 (40) 30024-30035
• 19 Chari R, Getz T, Nagy Jr B. et al. Protein kinase C[delta] differentially regulates platelet functional responses. Arterioscler Thromb Vasc Biol 2009; 29 (05) 699-705
• 20 Liverani E, Mondrinos MJ, Sun S, Kunapuli SP, Kilpatrick LE. Role of protein kinase C-delta in regulating platelet activation and platelet-leukocyte interaction during sepsis. PLoS One 2018; 13 (04) e0195379
• 21 Kostyak JC, Bhavanasi D, Liverani E, McKenzie SE, Kunapuli SP. Protein kinase C δ deficiency enhances megakaryopoiesis and recovery from thrombocytopenia. Arterioscler Thromb Vasc Biol 2014; 34 (12) 2579-2585
• 22 Kilpatrick LE, Sun S, Mackie D, Baik F, Li H, Korchak HM. Regulation of TNF mediated antiapoptotic signaling in human neutrophils: role of delta-PKC and ERK1/2. J Leukoc Biol 2006; 80 (06) 1512-1521
• 23 Kilpatrick LE, Standage SW, Li H. et al. Protection against sepsis-induced lung injury by selective inhibition of protein kinase C-δ (δ-PKC). J Leukoc Biol 2011; 89 (01) 3-10
• 24 Chen L, Hahn H, Wu G. et al. Opposing cardioprotective actions and parallel hypertrophic effects of delta PKC and epsilon PKC. Proc Natl Acad Sci U S A 2001; 98 (20) 11114-11119
• 25 Zaid Y, Marhoume F, Senhaji N. et al. Paraphenylene diamine exacerbates platelet aggregation and thrombus formation in response to a low dose of collagen. J Toxicol Sci 2016; 41 (01) 123-128
• 26 Caron A, Théorêt JF, Mousa SA, Merhi Y. Anti-platelet effects of GPIIb/IIIa and P-selectin antagonism, platelet activation, and binding to neutrophils. J Cardiovasc Pharmacol 2002; 40 (02) 296-306
• 27 Curtis MJ, Alexander S, Cirino G. et al. Experimental design and analysis and their reporting II: updated and simplified guidance for authors and peer reviewers. Br J Pharmacol 2018; 175 (07) 987-993
• 28 Zaid Y, Senhaji N, Naya A, Fadainia C, Kojok K. PKCs in thrombus formation. Pathol Biol (Paris) 2015; 63 (06) 268-271
• 29 Gawaz M, Langer H, May AE. Platelets in inflammation and atherogenesis. J Clin Invest 2005; 115 (12) 3378-3384
• 30 Totani L, Evangelista V. Platelet-leukocyte interactions in cardiovascular disease and beyond. Arterioscler Thromb Vasc Biol 2010; 30 (12) 2357-2361
• 31 Crosby D, Poole AW. Physical and functional interaction between protein kinase C delta and Fyn tyrosine kinase in human platelets. J Biol Chem 2003; 278 (27) 24533-24541
• 32 Pula G, Schuh K, Nakayama K, Nakayama KI, Walter U, Poole AW. PKCdelta regulates collagen-induced platelet aggregation through inhibition of VASP-mediated filopodia formation. Blood 2006; 108 (13) 4035-4044
• 33 Zaid Y, Senhaji N, Darif Y, Kojok K, Oudghiri M, Naya A. Distinctive roles of PKC delta isozyme in platelet function. Curr Res Transl Med 2016; 64 (03) 135-139
• 34 Murugappan S, Shankar H, Bhamidipati S, Dorsam RT, Jin J, Kunapuli SP. Molecular mechanism and functional implications of thrombin-mediated tyrosine phosphorylation of PKCdelta in platelets. Blood 2005; 106 (02) 550-557
• 35 Hall KJ, Jones ML, Poole AW. Coincident regulation of PKCdelta in human platelets by phosphorylation of Tyr311 and Tyr565 and phospholipase C signalling. Biochem J 2007; 406 (03) 501-509
• 36 Soroush F, Tang Y, Guglielmo K. et al. Protein kinase C-Delta (PKCδ) tyrosine phosphorylation is a critical regulator of neutrophil-endothelial cell interaction in inflammation. Shock 2019; 51 (05) 538-547