METHODS

Hearts were recovered postmortem from two patients who died eight and 52 weeks following PMR (of non-PMR related causes). Specimens were fixed in 4% formalin, embedded in paraffin, and 4 μm serial sections stained with haematoxylin. Sections were immunostained with monoclonal antibodies against α-smooth muscle actin (0.25 μg/ml), and polyclonal antibodies against von Willebrand factor (vWF)/factor VIII related antigen (1/200), and pan VEGF (2 μg/ml). Samples were also immunostained for CD45 and CD68 to assess the presence of lymphocytes and macrophages, respectively. Primary antibodies were detected with biotin-conjugated secondary antibodies and the ABC system (DAKO Ltd, Cambridge, UK). Antibodies were from DAKO except for the pan specific VEGF antibody that was from Santa Cruz Biotechnology (Santa Cruz, California, USA).

RESULTS

Both patients responded positively to PMR with apparent symptomatic improvement. Areas of fibrotic scarring were evident at eight and 52 weeks following PMR, that were not present in sections of the non-treated regions of left ventricle. These fibrotic zones contained a high density of microvessels as evidenced by positive staining for vWF (fig 1). A prominent feature in these areas was the presence of large apparently thin walled vessels. These were evident not only at eight weeks, but were particularly notable at 52 weeks post-PMR. Neovessels, including the abnormal large vessels, in PMR treated regions at week 8 and 52 weeks contained blood cells (fig 1), suggesting that they were perfused.

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Figure 1

Comparison of vessels in control and PMR treated myocardium. Sections of non-treated left ventricular anterior wall from eight week patient (A); non-treated area of left ventricular lateral wall from 52 week patient (B); PMR treated at eight weeks (C) and 52 weeks (D) immunostained for von Willebrand factor. Negative control without primary antibody is shown in D. Examples of established microvessels and equivalent sized PMR induced vessels are indicated by arrows and large diameter vessels by arrowheads. Asterisks indicate possible vessel fusions. Bar = 200 μm.

To investigate microvessel maturity, sections were probed for the smooth muscle cell marker, α-smooth muscle actin (fig 2A–C). Microvessels in non-treated left ventricular myocardium had prominent smooth muscle cell coverage (fig 2A), as did microvessels outside the areas of laser scarring in PMR treated ventricles (fig 2B,C). In contrast, many of the microvessels within the PMR induced fibrotic areas had little or no smooth muscle cell cover, either at eight or 52 weeks post-treatment (Fig 2B,C). There was no evidence of differences in lymphocyte infiltration associated with any areas of the PMR treated ventricles. Immunostaining with anti-CD68 antibodies demonstrated that macrophages were not present in either PMR treated or control sections. Using an antibody which recognises VEGF isoforms 121, 165, and 189, this angiogenic factor was detected in untreated areas of the heart as well as PMR treated regions (fig 2). In control regions of left ventricle, VEGF was detected in cardiac myocytes and smooth muscle cells surrounding microvessels. Areas of fibrosis lacked VEGF, and although smooth muscle cells surrounding microvessels within these regions stained positively for the growth factor, vessels lacking this investment were VEGF negative.

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Figure 2

Smooth muscle investment of vessels, presence of inflammatory cells and VEGF in control and PMR treated myocardium. Sections from non-treated areas (A) and PMR treated areas of myocardium at eight weeks (B) and 52 weeks (C) immunostained for von Willebrand factor (brown) and α-smooth muscle actin (red). VEGF (D, E). Negative control, primary antibody blocked with VEGF antigen peptide shown in inset (E). In panels A–C arrows indicate smooth muscle poor vessels and arrowheads indicate smooth muscle positive vessels. Bar = 100 μm.

DISCUSSION

This is the first report of sustained neovascularisation within the myocardium of human patients following PMR therapy. Clear evidence of increased numbers of neovessels was found not only shortly after PMR (eight weeks) but also longer term post-treatment (52 weeks). Notably the neovessels were abnormal. Specifically, many of these neovessels were smooth muscle cell negative and were large, dilated, thin walled vessels. Vessels lacking the regulatory influences on size and structure normally exerted by smooth muscle cells could remodel and expand to produce the abnormal vasculature observed. Restriction of the neovessels to fibrotic tissue, as well as their immature nature, probably limits the impact these vessels have on myocardial perfusion.

Blood vessels form initially as endothelial tubes, which then become invested with supporting smooth muscle cells during vessel maturation. Before acquisition of supporting cells, newly formed microvessels are unstable and, in the absence of survival factors such as angiogenic growth factors, undergo regression.⁴ The continued existence of apparently immature neovessels within PMR treated ventricles even at 52 weeks would indicate the presence of such angiogenic factors. A likely candidate for maintenance of the immature neovessels is VEGF, which was expressed in myocardial cells of both PMR treated and untreated areas of ventricles. Levels of this growth factor in the heart may be sufficient to support the immature vessels and allow them to remodel into the dilated, thin walled structures observed here, but not enough to initiate angiogenesis. It is a strong possibility, therefore, that other strategies to initiate neovascularisation in a similar myocardial environment in patients with chronic coronary disease, such as application of growth factors,⁵ may also result in formation and persistence of abnormal vessels. The present findings emphasise the need for a better understanding of mechanisms controlling formation and maturation of neovessels in the human heart in order to optimise revascularisation strategies, be they PMR based or involving direct delivery of angiogenic factors.