Acta Biomaterialia xxx (2014) xxx–xxx

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Brief communication

A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction Robert C. Rennert a,1, Kristine Rustad a,1, Kemal Levi a, Mark Harwood b, Michael Sorkin a, Victor W. Wong a, Amin Al-Ahmad b, Paul Zei b, Henry Hsia c, Ramin E. Beygui d, Linda Norton b, Paul Wang b, Geoffrey C. Gurtner a,⇑ a

Department of Surgery, Stanford University Medical Center, Stanford, CA, USA Division of Cardiovascular Medicine, Department of Internal Medicine, Stanford University Medical Center, Stanford, CA, USA c Division of Cardiology, Department of Internal Medicine, UCSF, San Francisco, CA, USA d Department of Cardiothoracic Surgery, Stanford University Medical Center, Stanford, CA, USA b

a r t i c l e

i n f o

Article history: Received 12 September 2013 Received in revised form 19 November 2013 Accepted 7 January 2014 Available online xxxx Keywords: Fibrosis Pacemaker Tensile test Stress analysis

a b s t r a c t The major risks of pacemaker and implantable cardioverter defibrillator extraction are attributable to the fibrotic tissue that encases them in situ, yet little is known about the cellular and functional properties of this response. In the present research, we performed a histological and mechanical analysis of human tissue collected from the lead–tissue interface to better understand this process and provide insights for the improvement of lead design and extraction. The lead–tissue interface consisted of a thin cellular layer underlying a smooth, acellular surface, followed by a circumferentially organized collagen-rich matrix. 51.8 ± 4.9% of cells were myofibroblasts via immunohistochemistry, with these cells displaying a similar circumferential organization. Upon mechanical testing, samples exhibited a triphasic force–displacement response consisting of a toe region during initial tensioning, a linear elastic region and a yield and failure region. Mean fracture load was 5.6 ± 2.1 N, and mean circumferential stress at failure was 9.5 ± 4.1 MPa. While the low cellularity and fibrotic composition of tissue observed herein is consistent with a foreign body reaction to an implanted material, the significant myofibroblast response provides a mechanical explanation for the contractile forces complicating extractions. Moreover, the tensile properties of this tissue suggest the feasibility of circumferential mechanical tissue disruption, similar to balloon angioplasty devices, as a novel approach to assist with lead extraction. Ó 2014 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

1. Introduction The number of pacemakers and implantable cardioverter defibrillators (ICDs) that are implanted each year continues to increase [1,2]. However, device-related complication rates are also rising [3], portending an increase in the rate of endovascular lead extraction procedures. While the indications for extraction have been established, and include systemic infections, pocket infections and lead malfunction [4], the potential for major cardiovascular injury or death with this procedure remains [5], despite the increased efficacy of laser-assisted techniques [6]. In fact, the recent multi-center Lead Extraction in the Contemporary Setting (LExICon) Study, which looked only at laser-assisted lead extraction, reported a procedural major adverse event rate of 1.4%, and an overall in-hospital mortality rate of 1.86% [7]. ⇑ Corresponding author. Tel.: +1 650 724 6672; fax: +1 650 724 9501. 1

E-mail address: [email protected] (G.C. Gurtner). These authors contributed equally to this work.

Interestingly, the major risks associated with lead extraction, including cardiac avulsions and vascular tears [7], can be directly linked to an imperfect separation of the lead from the fibrous tissue that binds it to the myocardium and venous endothelium. Moreover, this fibrotic response appears to be dynamic, as lead implantation time has been linked to an increase in complexity and overall failure of lead removal procedures [7,8], likely stemming from an increase in fibrosis along the lead body and electrode tip over time. Unfortunately, studies examining the in situ response to implantable pacemaker and ICD leads are rare [9,10], with relatively little known about the histological and mechanical features of the resulting tissue. A better understanding of this process could help improve the design of leads and extraction techniques, and further decrease the morbidity and mortality associated with this procedure. As such, this study aims to characterize the cellular and mechanical properties of human tissue collected from the lead surface upon removal of implantable devices, using stains, imaging

1742-7061/$ - see front matter Ó 2014 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.actbio.2014.01.008

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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R.C. Rennert et al. / Acta Biomaterialia xxx (2014) xxx–xxx

and mechanical analyses that have been established for the description of scarring and fibrosis in other organs.

2. Materials and methods 2.1. Extraction and collection of tissue All patients were consented as per the IRB protocol prior to their extraction procedure (IRB #19053). Procedures were performed in the operating room with electrophysiology and cardiac surgery physicians present. Pacemaker and/or ICD leads were extracted either by manual traction alone, or with a combination of Excimer laser sheaths (Spectranetics, Colorado Springs, CO) and manual traction. Upon completion of the procedure, all attached tissue was kept on the lead, and the lead and tissue were placed into 50 ml conical polypropylene tubes (BD Falcon, San Jose, CA) and put on ice. After collection, leads were promptly sent to the laboratory for analysis.

2.2. Histological analysis Tissue surrounding freshly extracted cardiac leads was carefully removed using microsurgical instruments, fixed in 4% paraformaldehyde and embedded in paraffin blocks, before being cut into 10 lm thick sections. After deparaffinization and rehydration, sections were stained with hematoxylin and eosin (H&E, Sigma–Aldrich, St Louis, MO), Masson’s trichrome (Sigma–Aldrich) or direct red 80 (picrosirius red, Sigma–Aldrich). Sections were also immunostained for type I and III collagens (primary 1:100, Abcam, Cambridge, MA; secondary 1:400 Alexa-Fluor 594 & 488, Invitrogen, Grand Island, NY), with nuclei stained with 40 ,6-diamidino2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA). Immunolabeling using a directly conjugated antibody against asmooth muscle actin (a-SMA) (Cy3, 1:250, Sigma–Aldrich, St Louis, MO), and for the intermediate filaments vimentin and desmin (primary 1:100, Abcam; secondary 1:400 Alexa-Fluor 594, Invitrogen) was similarly performed.

2.4. Tissue thickness quantification To quantify tissue thickness, four high-power images (400) were taken (at the 12, 3, 6 and 9 o’clock positions) of a single H&E stained tissue section per patient. Images were opened in AxioVision 4.8 software (Carl Zeiss, Inc, Oberkochen, Germany), and tissue thickness was measured once at each position (excluding irregular tissue appendages). Only patients with intact, circular tissue sections were included in average tissue thickness calculations. 2.5. Scanning electron microscopy (SEM) imaging For high-resolution SEM characterization of tissue, fresh tissue sections were cut to expose the tissue–lead interface and fixed in 4% paraformaldehyde, 2% gluteraldehyde and 0.1 M cacodylate buffer overnight. The tissue was then serially dehydrated in graded ethanol/water solutions, critically dried using a Tousimis Autosamdri-814 critical point dryer (Tousimis Research Corporation, Rockville, MD) and sputter-coated with a conductive layer of gold– palladium (AuPd). The samples were imaged using a Hitachi 3400 N VP scanning electron microscope (Hitachi High Technologies America, Inc., Schaumburg, Illinois) at the Stanford Cell Sciences Imaging Facility. 2.6. Mechanical testing Following technical refinements, selected tissue samples were immediately placed in phosphate-buffered saline upon collection, and stored at either –20 °C or 4 °C for mechanical testing. Tissue placed at –20 °C was analyzed within weeks of freezing, while tissue stored at 4 °C was analyzed within 72 h. The circumferential force–length relationships of samples were determined utilizing an adapted tensile ring methodology [11]. Briefly, circumferential samples were cut to create tissue rings with an approximate width of 1.5 mm. The rings were then loaded onto hooks of a uniaxial tensile testing apparatus (Bionix 200, MTS Systems Corporation, Eden Prairie, MN) equipped with a 44.48 N load cell and stretched until failure using a strain rate of 1% s1. The circumferential stress (rC) was calculated by dividing the load (L) at a given time point by two times the cross-sectional area (2⁄A) of the specimens at that given time point:

L 2A

2.3. Myofibroblast quantification

rC ¼

To quantify myofibroblasts, four high-power images (400) were taken (at the 12, 3, 6 and 9 o’clock positions) of a single tissue section per patient for both alpha-smooth muscle actin (a-SMA) and DAPI using an Axioplan 2 Imaging microscope (Carl Zeiss, Inc., Oberkochen, Germany), and overlayed in Photoshop CS5 (Adobe, San Jose, CA). Only patients with intact, circular tissue sections and positive staining were analyzed. Double positive and total cells were then counted for each picture by three independent investigators. Total a-SMA positive cell counts were averaged, normalized to total nuclei and reported as the mean percentage of aSMA positive nuclei per total cells. To ensure that vascular smooth muscle and endothelial cells (also stained by a-SMA) were excluded from this analysis, tissue blood vessel frequency and morphology were characterized on separate sections stained with H&E and directly conjugated antiisolectin (Alexa-Fluor 488, Griffonia simplicifolia, 1:200, Invitrogen). Cells exhibiting blood vessel morphology were found to be discrete and rare, and were excluded from myofibroblast cell counts. Expression levels of the intermediate filaments vimentin and desmin were also determined to confirm the myofibroblast phenotype.

Initial cross-sectional areas were calculated from baseline tissue width and thickness measurements, with cross-sectional areas at various levels of stretch determined as previously described based upon an assumption of isovolumic tissue deformation [11]. Circumferential strain (eC) was defined as the change in circumference/original circumference, and was determined to be the same as the diametric strain (strain based on diameter) as shown in the following equation (D = inner tissue ring diameter):

eC ¼

pðD þ DDÞ  DD DD ¼ : D pD

The circumferential tissue modulus was determined by plotting circumferential stress vs. circumferential strain, and calculating the slope of the line from the strain frame 1.5 to 2, as this initial linear portion of the curve was present in all samples. 2.7. Statistical analysis Quantitative data are expressed as mean ± standard error of the mean. Statistical significance across two groups was determined using a Student’s t-test, and across multiple groups using a one-

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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way ANOVA (SPSS Statistics v20.0, IBM, Armonk, NY). For all tests, a p-value  0.05 was considered statistically significant. 3. Results 3.1. Patient characteristics and tissue properties Over a 2 year period, 43 total leads were collected from 20 patients, with 28 leads (65%) having attached tissue. The average patient age at time of lead removal was 60.4 years, with the average time from lead implant being 5.2 years (Table 1). 15 of the 20 patients (75%) underwent lead removal as a result of infection, with nine pacemaker pocket infections (45%), four lead vegetations (20%) and two cases of bacteremia (10%). Outside of infectious complications, two patients underwent implant removal due to device erosion through tissue (10%) and one patient experienced a lead fracture (5%). The two remaining patients underwent lead removal as a result of patient pain or for lead management following factory recall. 3.2. Tissue histology and cellular composition The amount and thickness of tissue obtained from the leads varied across patients, with scar tissue often being found not only on the coiled surfaces and the lead tips, but also on the intravascularly located lead body (Fig. 1a and b). Leads without adherent tissue were excluded from further analysis. The average thickness of circumferential tissue samples was 458.6 ± 117.4 lm (n = 7 patients), with this variation likely stemming from the use of multiple extraction techniques (i.e. traction alone or laser-assisted). H&E, Masson’s trichrome and picrosirius red staining of tissue (n = 8 patients) showed a collagen-rich matrix with a low cellularity, and densely spaced collagen fibers demonstrating a circumferential organizational pattern (Fig. 1c and d, Supplemental Fig. 1a). Collagen organization and temporal dynamics were further characterized through picrosirius red staining, wherein thinner, less densely packed fibers tend to be yellowish-green, while thicker, more closely packed fibers are more likely to be orange-red [12,13]. Interestingly, this analysis revealed a trend toward smaller diameter fibers near the lead interface in the initial 7–8 months after implantation (Fig. 1d, left panels), suggestive of a predomi-

nant deposition of immature type III collagen known to occur in the early stages of wound healing [14,15]. Conversely, more uniform, thicker diameter collagen fibers were frequently observed in tissue samples obtained from leads that had been in place longer (Fig. 1d, right panels), consistent with normal scar remodeling and a transition to more organized and mature type I fibers. Collagen immunolabeling confirmed the predominance of type I collagen in more mature post-implantation fibrotic tissues (Fig. 1e); however, the persistence of lower levels of type III fibers in these samples is suggestive of an ongoing remodeling process. Immunohistochemical staining (n = 5 patients) revealed that 51.8 ± 4.9% of total cells expressed the myofibroblast marker aSMA, with many of these cells displaying a spindle-like morphology characteristic of myofibroblasts (Fig. 2a). Vimentin immunohistochemistry demonstrated a similar staining pattern as aSMA, while desmin staining was largely negative (Fig. 2b and c), with this a-SMA(+)/vimentin(+)/desmin() cell profile matching previous descriptions of myofibroblasts in pathologic fibrovascular and myocardial tissues [16,17]. These findings are consistent with a significant, sustained myofibroblast response on the level of other pathologic fibroses, such as hypertrophic scars [18], liver fibrosis [19] and idiopathic pulmonary fibrosis [20]. Additionally, a-SMA positive myofibroblasts displayed cytoplasmic extensions organized along the circumferential axis of the lead, suggestive of a potential role in the generation of contractile forces upon the implant. 3.3. SEM analysis SEM analysis (n = 3 patients) revealed that the lead–tissue interface was composed of a narrow layer of cells (likely monocytic based on morphology and presence of a foreign body) underlying a smooth, acellular surface, followed by a deeper, more fibrous layer with a primarily circumferential organization (Fig. 3a–c). Additionally, tissue ingrowth into the ICD coils was readily visualized (Fig. 3d), and found to occur as early as 6 months after implantation. 3.4. Mechanical properties Mechanical analysis of fibrotic tissue (n = 3 patients, 7 samples) generated force–displacement curves with three discrete regions:

Table 1 Patient and lead characteristics. Patient

Age (yrs)

Gender

Reason for Removal

Number of Leads

Lead Location

Implant Type

Time from Implant (months)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

57 65 53 60 75 67 91 48 67 63 67 54 65 68 78 18 45 78 47 41

F M F M M F M F F M M M M F M F F M M M

Pocket infection Pocket infection Lead vegetation Pocket infection Lead vegetation Pocket infection Bacteremia Lead fracture Patient pain Pocket infection Pocket infection Lead vegetation Pocket infection Lead vegetation Device erosion Device recall Device erosion Pocket infection Bacteremia Pocket infection

2 1 4 2 1 4 1 1 3 2 2 3 2 2 2 2 2 2 2 3

RA, RV RV RA (2), RV, LV RA, RV RV RA, RV(2), LV RV RA, RV RA, RV, LV RA, RV RA, RV RA, RV, LV RA, RV RA, RV RA, RV RA, RV RA, RV RA, RV RA, RV RA, RV(2)

P. D D P. D D D RD P D P. D P, D P. D P, D P RD P P.D P. 0 P P.D P. D

84 85 78,7 76 85 137, 88 6 10 11 106 43 67 224 84 44, 92 68 56 1.2 38 18

Patients with multiple time from implant dates reflect unique implantation procedures. RA, right atrium; RV, right ventricle; LV, left ventricle; P, pacer; D, defibrillator.

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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Fig. 1. Collection and characterization of lead tissue samples. (A) Schematic of pacemaker and defibrillator leads in situ. Green dots indicate commonly observed sites of fibrosis. (B) Gross appearance of leads illustrating the variation in fibrotic tissue adhesions across samples. (i–iii) RV defibrillator lead explanted after 85 months with fibrosis at the lead tip and along the proximal (intravascular) lead body. (iv) RV pacer explanted after 167 months with large fibrotic adhesions at the lead tip. (v) RV pacer explanted after 6 months with minimal adherent tissue. (C) Low- and high-power representative images of tissue following H&E (left panels) and Masson’s trichrome (right panels) staining, demonstrating the low cellularity and high collagen content of samples. (D) Bright-field microscopy of picrosirius red stained samples with birefringence specific for collagen (larger collagen fibers tend to be orange-red, while thinner ones are generally yellowish-green) redemonstrates the circumferential organization and suggests a progressive collagen remodeling. In the left panels, smaller diameter fibers are visualized near the lead interface, which was removed only 7 months after implantation. In the right panels, the lead interface demonstrates the presence of a more mature scar 76 months following implantation, with a uniform, thicker diameter collagen composition. (E) Representative immunohistochemical staining for collagen type I (red) and III (green) (blue – nuclear marker DAPI) illustrating the heterogeneous fiber composition of surrounding fibrotic tissues (RV defibrillator sample explanted after 85 months) (scale bar = 100 lm in all images).

an initial toe region when the tissue became fully tensioned, a linear region during which the tissue displayed elastic properties and a yield and failure region where initial tearing began and continued to a maximum force causing rupture (Fig. 4a–c). The mean fracture load across samples was 5.6 ± 2.1 N (range 3.1–9.5 N), while the mean calculated circumferential stress at which the tissue failed was 9.5 ± 4.1 MPa (range 4.5 to 16.9 MPa) (Fig. 4d). For comparison, these values are similar to those reported for other semi-elastic human tissues, such as uninjured skin and cardiac tissue

[21,22], but 1000 times lower than the forces required for the fracture of long bones [23]. Additionally, the mean fracture strain across samples was 3.4 ± 0.6 (range 2.6–4.4), while the circumferential tissue modulus (based upon a strain frame of 1.5 to 2) was 3.4 ± 0.5 MPa (range 2.8–4.1 MPa) (Fig. 4d). Of note is that patient origin and storage conditions prior to testing showed no significant impact on mechanical properties (p = 0.38 and 0.42 for fracture load, p = 0.47 and 0.61 for fracture stress, p = 0.75 and 0.82 for fracture strain and p = 0.10 and 0.16 for the tissue modulus).

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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Fig. 2. Myofibroblasts are a large component of the fibrotic response to implanted leads. (A) Immunohistochemical staining and quantification of the myofibroblast marker aSMA (red; blue – nuclear stain DAPI). Approximately 50% of total cells are aSMA (+), with many displaying a spindle cell morphology characteristic of myofibroblasts (white arrowheads). (B, C) Fibrotic tissues were also positive for the intermediate filament vimentin, and largely negative for desmin, consistent with a myofibroblast phenotype (scale bar = 100 lm in all images).

Fig. 3. SEM visualization of scar tissue. (A, B) Low, medium and high magnification SEM images of collected tissue illustrating a narrow layer of cells (black arrowheads) underlying a smooth lead–tissue interface (white arrowheads), followed by a deeper, circumferentially organized fibrous layer (brown arrowheads). (C) High magnification SEM visualization of the smooth lead–tissue interface. (D) SEM visualization of tissue ingrowth into lead coils (white arrowheads).

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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Fig. 4. Analysis of mechanical properties of scar tissue. (A) Depiction of experimental setup for determination of the circumferential mechanical properties of tissue. The circumferential stress (rC) was calculated by dividing the load (L) at a given time point by two times the cross-sectional area (2 * A) of the specimens at that given time point  rC ¼ 2AL . Circumferential strain (eC) was defined as the change in circumference/original circumference, and was determined to be the same as the diametric strain   pðDþDDÞDD eC ¼ ¼ DDD , where D = the inner tissue ring diameter. (B) Representative images from mechanical analysis depicting a sample pre-load (left image) and beginning pD to fail (right image). Failure occurred at approximately the midpoint of all tissue samples (white arrow). (C) Force–displacement curves of samples, illustrating the toe, linear and yield/failure regions. Average circumferential stress is shown in red. (D) Sample cross-sectional areas, fracture loads and calculated circumferential fracture stress/strain and circumferential tissue modulus values. The circumferential tissue modulus was defined as the slope of the circumferential stress vs. circumferential strain curve from the strain frame of 1.5 to 2, as this initial linear portion was present in all samples.

4. Discussion In designing this study, we hypothesized that a better characterization of the fibrotic response to lead implants could be used to improve both the design of leads and extraction techniques. The discussion of our findings is therefore focused around this hypothesis. Overall, the fibrosis-dominated response to lead implants characterized herein is consistent with a foreign body reaction to an implanted material. This response is an attempt by the body to isolate the implant from the local environment, and is typically characterized by an acute inflammatory cell infiltrate that progresses to a macrophage-dominated chronic inflammatory phase, and ultimately results in fibrotic capsule formation following fibroblast recruitment and proliferation [24,25]. The foreign body response is not fixed, however, as the relative level of fibrosis and inflammation within reactive tissue can be influenced by both surface characteristics of the implant, as well as the cellular composition of affected tissue [25]. For example, implants with high surface-to-volume exteriors can promote an increased macrophage response characterized by the formation of foreign body giant cells at the implant surface, which can ultimately contribute to device failure from persistent oxidative stress [24,25]. Exerting a similar modulatory influence, permanent and stable cell types (e.g. cardiomyocytes and endothelial cells) are thought to have an increased propensity for fibrosis following

foreign body insult, compared to the more regenerative response of cell populations with a higher replicative capacity (e.g. epithelial cells) [25]. Interestingly, mechanotransduction pathways can also affect both inflammation and fibrosis [26,27], and are likely contributors to the foreign body response in this scenario as the introduction of a static implant to the dynamic environment of the cardiovascular system undoubtedly creates mechanical force perturbations. While the presence of an infection (the reason for removal in the majority of leads) may also influence the fibrotic response, the long-term asymptomatic period of lead functionality prior to removal, as well as the presence of a similar fibrotic response in non-infected leads, suggests this is not a main contributor to the formation of fibrosis. Interpreted in this light, the low cellularity and high collagen content of the fibrotic tissue in this study is most likely due to both the low regenerative capacity of the cell types in contact with the cardiac implants and advancements in implantable material design specifically made to limit their resulting macrophage response [24,25,28]. While these findings are largely consistent with the evolving definition of biocompatibility as it relates to long term implantable devices, namely the ability to perform a desired function without eliciting an undesired local or systemic effect [29], the potential for fibrosis-related mitigation of electrical impulses at the lead tip, a focal point for adhesions in this study, remains a concern. In fact, the dynamic nature of this response is illustrated by the eventual formation of a smooth surfaced capsule at the

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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implant–tissue interface seen with SEM, as well as the remodeling of collagen fibers suggested by immunohistochemical and picrosirius red staining, which is similar to that observed in injured skin and myocardium [12]. Some level of fibrosis is nonetheless desirable to maintain lead stability following implantation. However, given the strikingly high coefficients of friction encountered during specimen collection, we hypothesized that either tissue ingrowth occurred along the length of the lead, or contractile forces must be present, such as those known to occur in injured skin and cardiac tissue as a result of myofibroblast infiltration and organization [16,30,31]. In light of the absence of macro- or microscopic tissue ingrowth along the lead bodies, we reasoned that the main force resisting extraction was generated as part of the normal fibrotic response to an implant, potentially exacerbated by the robust mechanical stimulation of the intravascular environment. It was therefore not surprising that approximately half of all nucleated cells in the lead–tissue samples were positive for the myofibroblast marker a-SMA, with many of these cells displaying cytoplasmic extensions consistent with the exertion of circumferential contractile forces. Although myofibroblasts display variability in cytoskeletal patterning that make their definitive immunohistochemical identification challenging [32], the observed a-SMA(+)/vimentin(+)/desmin() profile is consistent with previous descriptions of myofibroblasts within pathologic fibrovascular and myocardial tissues [16,17]. Moreover, the collagen-rich and relatively acellular nature of the tissue samples (as characterized by histology and electron microscopy) makes the presence of other potentially similarly staining cell types, such as vascular smooth muscle and mature cardiomyocytes, unlikely. While typically absent in normal tissues [33], myofibroblasts are known to rapidly populate sites of injury in response to proinflammatory cytokines [34]. These cells likely originate from resident fibroblasts, as well as non-resident cells, such as monocytes and endothelial precursor cells, and following recruitment, provide mechanical tension to the remodeling matrix by anchoring and contracting [34]. Upon completion of normal repair processes, myofibroblasts typically undergo apoptosis [30]; however, they can persist under certain pathological conditions. Specifically, myofibroblasts have been shown to account for up to 65% of the fibroblast-like cells within hypertrophic scars [18], and more than 50% of all cells within the scar tissue of advanced stage fibrotic livers [19]. While not quantified, similarly high levels of myofibroblasts are also seen within the fibroproliferative clusters of idiopathic pulmonary fibrosis [20], as well as in the capsular response to foreign body implants [35]. Interestingly, cardiac tissue also displays a strong, sustained myofibroblast response following injury, with high levels of myofibroblasts organized along stress lines seen decades after ischemic events [16]. The prolonged myofibroblast response observed in this study is therefore not unique, but almost certainly contributes to the high lead–tissue coefficient of friction through the generation of circumferential forces. Most importantly, the identification of a large myofibroblast response in this setting suggests that a more controlled regulation of contractile forces on the implant is possible, especially given recent advancements in our ability to modulate myofibroblast differentiation [36]. The mechanical analyses performed in this study were designed to provide another dimension to the characterization of collected tissues. In planning these studies, the most clinically relevant mechanical aspect was determined to be circumferential tensile strength and failure load, as these properties dictate the ease of radial tissue disruption and ability to separate the surrounding fibrosis from the implant surface. We therefore calculated the circumferential mechanical properties of this tissue utilizing the experimental setup described in Fig. 4a, ignoring the likely

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variable elastic properties when stressed in other directions [22,37,38]. Not surprisingly, the collected tissue displayed mechanical properties consistent with other biomaterials. The toe, linear and failure regions observed on our stress–strain curves have been previously described for other tissue types, including skin and normal cardiac tissue [21,22,37,39]. This triphasic behavior has been ascribed largely to the collagen fibers within a given tissue, and corresponds to the sequential progression of collagen uncrimpring, stretching of collagen triple helices or cross-links between helices and ultimately disruption of the fibril structure that occurs with an increasing load [37,40–43]. While direct comparisons of the failure load and circumferential stress reported herein to other tissue types is difficult due to variations in experimental technique across studies, the validity of our findings is supported by reports of comparable failure loads of 4.9 N for normal cardiac tissue [21]. Similarly, our data are within the wide range (1–32 MPa) of previously reported tensile strength values for intact skin [22]. The narrow range of the circumferential tissue modulus is also supportive of the reproducibility of the experimental system, as well as a similarity in stiffness across individual samples. Extrapolating this mechanical data to the complicating nature of lead–tissue adhesions upon extraction, the development of leads with an expandable exterior, similar to balloon angioplasty devices, could theoretically be used to disrupt the surrounding fibrotic tissue and facilitate safe lead removal. Supporting the feasibility of this approach, the stress–displacement curves generated herein are likely representative of the maximum tensile properties of this fibrotic response, as these mechanical analyses were performed on tissue collected from leads with relatively long implantation times. Moreover, the forces necessary to produce sub-failure expansile stress within our samples, the delivery of which in situ would allow lead removal with minimial associated risk of vessel rupture, are achievable with current angioplasty balloon technology [44]. For rapid clinical translation, this methodology could even be combined with existing extraction techniques to accelerate the lead removal process, as the simultaneous or segmental expansion of a coating balloon on the non-conductive portions of implanted leads would likely be faster than the progressive tissue disruption of cutting sheaths along the entire lead length, with this more time-intensive process reserved exclusively for conductive portions of the lead that would not be amenable to an exterior coating. Throughout this study, quantitative correlation of sample properties to clinical data was dictated by the variable amount of tissue collected per lead. For example, while we believe that maturation of the fibrotic response suggested by the immunohistochemical and picrosirius red staining is almost certainly associated with an increase in tensile strength due to the known influence of collagen fibril diameter on the strain response to loading [45], the limited number of samples and patient implant times available for mechanical testing made a definitive analysis of the effect of time on fibrotic tissue strength outside the scope of this study. Nonetheless, the histological and mechanical data presented herein provide a novel, clinically relevant characterization of the tissue response to implants within the cardiovascular system, as well as feasibility data for researchers seeking to use mechanical scar disruption for the improvement of lead extraction techniques.

5. Conclusions Characterization of the tissue surrounding implanted cardiac pacemaker and ICD leads reveals a mostly low cellularity fibrotic response, with densely packed collagen fibers organized in a

Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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circumferential pattern around the implant. A sustained myofibroblast response is also present and similarly organized along the circumferential axis, likely providing circumferential contractile forces complicating lead removal. The mechanical properties of this tissue are similar to those reported for skin and uninjured myocardium, suggesting the possibility of novel, mechanically based extraction techniques. Financial disclosure The authors have no potential conflicts of interest, affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed herein. Acknowledgements The authors would like to thank Yujin Park for her assistance with tissue processing and staining, as well as Lydia-Marie Joubert at the Stanford Cell Imaging Facility for her assistance with electron microscopy. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.actbio. 2014.01.008. Appendix B. Figures with essential color discrimination Certain figures in this article, particularly Figures 1, 2, and 4, are difficult to interpret in black and white. The full color images can be found in the on-line version, at http://dx.doi.org/10.1016/ j.actbio.2014.01.008 References [1] Kurtz SM, Ochoa JA, Lau E, Shkolnikov Y, Pavri BB, Frisch D, et al. Implantation trends and patient profiles for pacemakers and implantable cardioverter defibrillators in the United States: 1993–2006. Pacing Clin Electrophysiol 2010;33(6):705–11. [2] Uslan DZ, Tleyjeh IM, Baddour LM, Friedman PA, Jenkins SM, St Sauver JL, et al. Temporal trends in permanent pacemaker implantation: a population-based study. Am Heart J 2008;155(5):896–903. [3] Voigt A, Shalaby A, Saba S. Rising rates of cardiac rhythm management device infections in the United States: 1996 through 2003. J Am Coll Cardiol 2006;48(3):590–1. [4] Wilkoff BL, Love CJ, Byrd CL, Bongiorni MG, Carrillo RG, Crossley 3rd GH, et al. Transvenous lead extraction: heart rhythm society expert consensus on facilities, training, indications, and patient management. Heart Rhythm 2009;6(7):1085–104. [5] Gaca JG, Lima B, Milano CA, Lin SS, Davis RD, Lowe JE, et al. Laser-assisted extraction of pacemaker and defibrillator leads: the role of the cardiac surgeon. Ann Thorac Surg 2009;87(5):1446–50. discussion 1450–1441. [6] Wilkoff BL, Byrd CL, Love CJ, Hayes DL, Sellers TD, Schaerf R, et al. Pacemaker lead extraction with the laser sheath: results of the pacing lead extraction with the excimer sheath (PLEXES) trial. J Am Coll Cardiol 1999;33(6):1671–6. [7] Wazni O, Epstein LM, Carrillo RG, Love C, Adler SW, Riggio DW, et al. Lead extraction in the contemporary setting: the LExICon study: an observational retrospective study of consecutive laser lead extractions. J Am Coll Cardiol 2010;55(6):579–86. [8] Bracke F, Meijer A, Van Gelder B. Extraction of pacemaker and implantable cardioverter defibrillator leads: patient and lead characteristics in relation to the requirement of extraction tools. Pacing Clin Electrophysiol 2002;25(7):1037–40. [9] Epstein AE, Kay GN, Plumb VJ, Dailey SM, Anderson PG. Gross and microscopic pathological changes associated with nonthoracotomy implantable defibrillator leads. Circulation 1998;98(15):1517–24. [10] Candinas R, Duru F, Schneider J, Luscher TF, Stokes K. Postmortem analysis of encapsulation around long-term ventricular endocardial pacing leads. Mayo Clin Proc 1999;74(2):120–5.

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Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

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Please cite this article in press as: Rennert RC et al. A histological and mechanical analysis of the cardiac lead–tissue interface: implications for lead extraction. Acta Biomater (2014), http://dx.doi.org/10.1016/j.actbio.2014.01.008

A histological and mechanical analysis of the cardiac lead-tissue interface: implications for lead extraction.

The major risks of pacemaker and implantable cardioverter defibrillator extraction are attributable to the fibrotic tissue that encases them in situ, ...
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