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Am J Physiol Lung Cell Mol Physiol 292: L742-L747, 2007. First published November 22, 2006; doi:10.1152/ajplung.00064.2006
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Vascular endothelial growth factor accelerates compensatory lung growth after unilateral pneumonectomy

Maromi K. Sakurai,1,2 Sang Lee,1,2 Danielle A. Arsenault,1,2 Vania Nose,3 Jay M. Wilson,1 John V. Heymach,1,2 and Mark Puder1,2

1Department of Surgery, and 2Vasculary Biology Program, Children's Hospital Boston, Harvard Medical School, Boston; and 3Department of Pathology, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts

Submitted 20 February 2006 ; accepted in final form 17 November 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We hypothesize that compensatory lung growth after unilateral pneumonectomy in a murine model is, in part, angiogenesis dependent and can be altered using angiogenic agents, possibly through regulation of endothelial cell proliferation and apoptosis. Left pneumonectomy was performed in mice. Mice were then treated with proangiogenic factors [vascular endothelial growth factor (VEGF); basic fibroblast growth factor (bFGF)], VEGF receptor antibodies (MF-1, DC101), and VEGF receptor small molecule chemical inhibitors. Lung volume and mass were measured. The lungs were analyzed using immunohistochemistry by CD31 staining, terminal deoxynucleotidyl transferase biotin-dUTP nick end labeling, type II pneumocytes staining, and proliferating cell nuclear antigen. Compensatory lung growth was complete by postoperative day 10 and was associated with diffuse apoptosis of endothelial cells and pneumocytes. This process was accelerated by VEGF, such that growth was complete by postoperative day 4 with similar associated apoptosis. bFGF had no effect on lung growth. MF-1 and DC101 had no effect. The VEGF receptor small molecule chemical inhibitors also had no effect. VEGF, but not bFGF, accelerates growth. VEGF receptor inhibitors do not block growth, suggesting that other proangiogenic factors play a role or can compensate for VEGF receptor blockade. Diffuse apoptosis, endothelial cell and pneumocyte, occurs at cessation of both normal compensatory and VEGF-accelerated growth. Angiogenesis modulators may control growth via regulation of endothelial cell proliferation and apoptosis, although the exact relationship between endothelial cells and pneumocytes has yet to be determined. The fact that bFGF did not accelerate growth in our model when it did accelerate regeneration in the liver model suggests that angiogenesis during organ regeneration is regulated in an organ-specific manner.

basic fibroblast growth factor; pneumocytes; lung hypoplasia

ANGIOGENESIS IS IMPORTANT in both pathological and physiological processes. Angiogenesis-dependent mechanisms, such as wound healing and embryonic organ development, have been under extensive investigation. Most noteworthy of these has been the well-established dependence of tumor tissue on angiogenesis (6). Tumors cannot survive or grow without adequate nutrients and oxygen, which are made available by blood vessels. Without appropriate sustenance, tumor cells die via apoptosis or necrosis (13). Tumors secrete various angiogenic factors to recruit new blood vessels, a process that is critical for tumor growth (7, 11). The physiological equivalents to tumor growth are de novo tissue growth (embryogenesis) and tissue regeneration. Angiogenesis is presumably a critical factor in these processes as well. Support for the role of angiogenesis in the development of lung tissue can be found in studies showing that the anti-angiogenic agents fumagillin, thalidomide, and SU-5416 attenuate vascular growth and reduce alveolarization in developing rat lung (15). Additionally, it has been shown that compensatory vascular growth in lung tissue is proportional to the amount of tissue removed during pneumonectomy (21). Taken together, this research suggests a significant role for angiogenesis in both the normal development and compensatory growth of the lung.

Our laboratory previously demonstrated that liver regeneration after partial hepatectomy is dependent on angiogenesis, and furthermore, that this process can be altered using angiogenesis modulators (10). The proangiogenic factor, basic fibroblast growth factor (bFGF), accelerated liver regeneration, whereas angiogenesis inhibition via TNP-470, a synthetic analog of the Aspergillus-derived antibiotic fumagillin and known inhibitor of endothelial cell proliferation and migration, restricted growth (9, 14, 32). It is now clear that there is a delicate regulatory interplay between tissue and vasculature, or more specifically endothelial cells, which is mediated by various growth factors (4, 20, 29, 30, 33, 36).

Vascular endothelial growth factor (VEGF) is another well-known proangiogenic factor which is endogenously produced and is important in the normal vascular development of numerous organ systems. In the pulmonary system, angiogenesis is of particular importance because the blood-air interface is the sole source of oxygen delivery to the body. VEGF has been shown to be deposited in the subepithelial matrix at the leading edges of branching airways where it stimulates angiogenesis (12). It has also been demonstrated that during pulmonary distention, a stimulus for alveolarization (capillary and alveolar growth) upregulated VEGF mRNA and led to the hypothesis that angiogenesis is a rate-limiting factor in this process (25). Additionally, lung tissue appears to self-regulate its own development with respect to vasculature in that type 2 pneumocytes and epithelial cells express both VEGF and VEGF receptors (17, 19). Similar to the liver, we see evidence for a tissue-vasculature relationship that may regulate tissue growth. In a series of experiments, Compernolle et al. (5) showed that HIF-2{alpha}, one of the hypoxia-inducible transcription factors of VEGF, was critical in the normal development of murine lungs and that exogenous VEGF administration was able to rescue premature animals from respiratory distress syndrome by improving aeration, increasing surfactant production, and improving mortality.

There are several models of compensatory lung growth after unilateral pneumonectomy. In rodents, left pneumonectomy elicits compensatory growth of the remaining right lung to total lung mass within 10 days (3). However, it does not exceed original organ size, similar to liver regeneration after partial hepatectomy. The mechanism controlling initiation, propagation, and termination of compensatory growth remains unclear. In addition, the relationship between pneumocytes and endothelial cells in growing lung tissue is unknown.

It is our hypothesis that compensatory lung growth after unilateral pneumonectomy in a murine model is, in part, angiogenesis dependent and can be altered with angiogenesis modulators, possibly through regulation of endothelial cell proliferation and apoptosis.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Surgical procedure. Disease-free 7- to 8-wk-old C57BL/6 male mice (Jackson Laboratories, Bar Harbor, ME), weighing ~25 g each, were anesthetized with 120–400 mg/kg Avertin (Sigma, St. Louis, MO) via intraperitoneal injection. Trans-oral intubation, thoracotomy, and excision of the left lung were performed as described previously (31, 38). Briefly, trans-oral intubation was performed using a 2.7-mm zero degree rigid endoscope (Karl Storz, Charlton, MA) and an 18-gauge angiocatheter (Edwards Lifesciences, Irvine, CA) as an endotracheal tube. Animals were ventilated with room air at 70 breaths/min using a rodent ventilator (HSE-HA Minivent, Harvard Apparatus, Holliston, MA). An anterior midaxillary incision was made, followed by a thoracotomy incision at the fifth intercostal space. The hilum of the left lung was then ligated with 5–0 silk suture (Ethicon, Somerville, NJ), and the lung was removed. Animals were then administered a 5-ml normal saline bolus subcutaneously and allowed to recover under warming lamps. The sham-operated mice underwent a simple left thoracotomy but the left lung was neither ligated nor removed. All animal experiments were done in accordance with National Institutes of Health guidelines as dictated by the Animal Care Facility at Children's Hospital Boston.

Lung harvest and fixation. Mice were weighed and euthanized on postoperative days 4, 8, 10, and 12. The remaining right lung was removed with the tracheo-bronchial tree intact and infused with 10% formalin via angiocatheter intubation of the trachea, at a pressure of 25 cmH2O, to fix the alveolar space and maintain pulmonary architecture (8). The lung was ligated at the hilum and the trachea was removed. Lung volumes were determined using a volume displacement method (34). The tissue sample was then immersed in 10% formalin at 4°C overnight. The next day, the sample was washed in PBS three times and immersed in 70% ethanol (EtOH) in PBS. The lungs were then sectioned in paraffin before immunohistochemical analysis.

Morphometry. Hemotoxylin and eosin (H&E)-stained lung sections were used for morphometric analysis. Lung morphometry was determined using a three-level sampling and point counting technique (18). Measured parameters were total lung volume, total air space volume, total alveolar tissue volume, and total alveolar surface density.

Angiogenesis modulator administration. Two angiogenesis enhancers were administered. VEGF-164 (National Cancer Institute, Biological Resources Branch, Frederick, MD), an endothelial mitogen, was administered at 0.5 mg/kg ip daily. bFGF (National Cancer Institute, Biological Resources Branch), a stimulator of endothelial cell proliferation and migration, was administered at 1 mg/kg ip daily. Mice treated with VEGF were killed on postoperative days 4 and 10, and those treated with bFGF were killed on postoperative day 4.

Four angiogenesis inhibitors were administered. MF-1, a selective VEGF receptor-1 (anti-VEGFR-1) monoclonal antibody (Imclone Systems, New York, NY), was administered at 32 mg/kg ip every third day. DC101, a selective VEGFR-2 (anti-VEGFR-2) monoclonal antibody (Imclone Systems), was administered at 32 mg/kg ip every third day. MF-1 and DC101 were also administered as a combination therapy at the above dosing schedule. SU11248, a selective VEGF receptor-2 small molecule chemical inhibitor (Pfizer, New York, NY), was administered at 40 mg·kg–1·day–1 by oral gavage daily. In addition, a selective VEGF receptor-1 small molecule inhibitor was administered at 100 mg·kg–1·day–1 by oral gavage daily. Mice treated with angiogenesis inhibitors were killed on postoperative day 10.

Immunohistochemistry. Paraffin lung sections were rehydrated with sequential xylene and EtOH washes. Sections were subjected to xylene immersion for 2 min twice, 100% EtOH for 3 min, 95% EtOH for 3 min, and then 70% EtOH for 3 min. Next, epitope retrieval was achieved by incubating sections with Proteinase K (20 µg/ml) in a humidified chamber at 37°C for 20 min, followed by a cool down period to room temperature for 20 min. Sections were then washed three times with PBS + 0.3% Triton X-100. Serum blocking was then performed by incubating sections in 5% normal goat serum (Jackson ImmunoResearch, West Grove, PA) in PBS + 0.3% Triton X-100 + 0.01% bovine serum albumin + 0.01% Thimerosal at 37°C for 30 min. Then, they were again washed three times with PBS + 0.3% Triton X-100. Primary antibody incubation with rat anti-mouse CD31 antibody (BD Biosciences Pharmingen, San Diego, CA) was diluted to 1:50 in PBS + 0.3% Triton X-100 + 0.01% bovine serum albumin + 0.01% Thimerosal and incubated overnight at room temperature. Samples were then washed three times with PBS + 0.3% Triton X-100. Secondary antibody incubation was then done with goat anti-rat IgG Alexa Fluor 594 (Molecular Probes, Invitrogen, Carlsbad, CA) diluted to 1:500 for 2 h at room temperature. Sections were washed three times with PBS + 0.3% Triton X-100.

After the last wash step in the CD31 staining protocol, the first step in the terminal deoxynucleotidyl transferase biotin-dUTP nick end labeling (TUNEL) staining protocol was performed using the DeadEnd Fluorometric TUNEL System (Promega, Madison, WI). Briefly, sections were covered with equilibration buffer provided in the kit for 5 to 10 min at room temperature. Next, rTDT enzyme mixture (45-µl equilibration buffer, 5-µl nucleotide mixture, 1 µl rTDT enzyme) was added to each section and incubated in a humidified chamber at 37°C for 60 min. Sections were immersed in 2 x SSC, in a coplin jar at room temperature for 15 min to terminate the reaction. Sections were washed for 5 min three times with PBS + 0.1% Triton X-100 + bovine serum albumin (5 mg/ml) and then washed once with PBS. Next, the sections are covered with Hoechst 33258 (1 µg/ml) nuclear stain (BisBenzimidine, Sigma) for 5 s. The stain was aspirated, and two drops of Gel Mount Aqueous Mounting Medium (Sigma) were applied followed by a coverslip. Clear nail polish was applied to the edges of the coverslip and allowed to dry.

ProSPC primary antibody specific for type II pneumocytes (Chemicon International, Temecula, CA) was also used as detailed above, substituting for CD31 primary antibody where noted. The secondary antibody was Cy3-conjugated AffiniPure goat anti-rabbit IgG (1:200 dilution; Jackson ImmunoResearch, West Grove, PA).

Proliferation assay. Proliferation was measured in both control and VEGF-treated groups, as previously described (10). Specifically, cell proliferation was assessed by proliferating cell nuclear antigen (PCNA) immunostaining. After deparaffinization and rehydration, lung sections were incubated with Citra buffer (Dako, Denmark) for 10 min at 90°C. Nonspecific binding sites were then blocked for 30 min at room temperature with TNB blocking buffer (New England Nuclear Life Sciences, Boston, MA). Primary PCNA anti-serum 1:150 (Signet Laboratories, Dedham, MA) was added to the sections overnight at 4°C. Secondary goat anti-mouse IgG 1:400 antibody was then added for 30 min at room temperature. Next, the slides were incubated with streptavidin alkaline phosphatase (AP) solution (1:100), followed by biotinyl tyramide amplification diluent (1:50). After repeat incubation with AP solution (1:100) for 30 min at room temperature, the chromagen New Fuchsin (Biogenex, San Ramon, CA) was added. The slides were counterstained with Gill's hematoxylin. After all incubations, except for the TNB blocking step, the sections were washed three times with TNT wash buffer (New England Nuclear Life Sciences). The TNB buffer, TNT buffer, biotinylated-tyramide solution, amplification diluent, and streptavidin-AP were used according to the instructions of the New England Nuclear TSA indirect kit (catalogue no. NEL 700A, New England Nuclear Life Sciences).

Confocal microscopy. All stained sections were analyzed using the Leica TCS SP2 confocal microscope and its associated software. Images were optimized for Z-position, gain, O/U flow, and laser intensity. Ten high-powered fields (x63) were selected at random and counted by a blinded observer for positive TUNEL staining.

Vascular density. Paraffin-sectioned lungs were stained for CD31 as previously described in this section. However, instead of a fluorescent secondary antibody, a chromagen-based secondary antibody was used for visualization under light microscopy. Fifteen high-powered fields (x100) from three different animals were counted for each section in a blinded fashion. One high-powered field corresponded to ~100 alveoli. Means were analyzed for statistical significance.

Statistical analysis. Statistical analysis for the lung morphometry data was performed using the Student's two-tailed, unpaired t-test for comparisons between groups. Differences were considered significant when P < 0.05. Raw counts from the immunohistochemistry data were subjected to Mann-Whitney rank sum test, using the Sigma Stat program (SPSS, Chicago, IL).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Compensatory lung growth. Following removal of the left lung, the right lung grew beyond its original size to compensate for that loss of lung mass. Compensatory lung growth was measured as a function of lung (volume and mass) to animal weight ratio vs. time. The original lung weight to body weight ratio was 5.6 ± 0.3 (x 10–3 g/g) and the original lung volume to body weight ratio was 41.8 ± 1.7 (x 10–3 ml/g). By postoperative day 8, complete compensatory lung growth was achieved (Figs. 1 and 2). Morphometric analysis revealed no significant difference in alveolar surface density, volume of respiratory regions, and volume of respiratory airspaces between sham-operated and pneumonectomy animals (data not shown).


Figure 1
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  		Fig. 1. Lung volume to body weight ratio (LV:BW) on postoperative day 10 after pneumonectomy. Error bars represent SE.

 

Figure 2
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  		Fig. 2. Time course of lung mass to body weight ratio (LM:BW) after pneumonectomy. Error bars represent SE.

 
Effects of angiogenesis modulators on compensatory lung growth. VEGF-treated mice showed complete compensatory lung growth at postoperative day 4, as measured by lung volume:weight and shown as percent compensatory growth (Fig. 3). VEGF accelerated compensatory lung growth from 8–10 to 4 days (P < 0.001). Morphometric analysis between control and VEGF-treated animals showed no significant difference in alveolar surface density, volume of respiratory regions, and volume of respiratory airspaces (data not shown). bFGF did not accelerate compensatory growth, and there was in fact a trend toward reduced growth, although this did not achieve statistical significance (P = 0.6; Fig. 3).


Figure 3
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  		Fig. 3. Percent compensatory lung growth by LV:BW ratio on postoperative day 4 after pneumonectomy (PTX) ± VEGF and basic fibroblast growth factor (bFGF). Error bars represent SE.

 
Neither MF-1 (anti-VEGFR-1) nor DC101 (anti-VEGFR-2) had any effect on compensatory lung growth (P = 0.558). Furthermore, the administration of MF-1 plus DC101 concurrently did not have any effect (P = 0.446; Fig. 4). In addition, the small molecule chemical inhibitors had no effect on compensatory lung growth (data not shown).


Figure 4
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  		Fig. 4. Percent compensatory lung growth by LV:BW ratio on postoperative day 10 after PTX ± DC101, MF-1, and DC101+MF-1. Error bars represent SE.

 
Immunohistochemistry. In untreated animals that underwent pneumonectomy, there was a significant increase in TUNEL-positive cells on postoperative day 10 compared with postoperative day 4 (P < 0.001). In VEGF-treated animals, there was a significant increase in TUNEL-positive cells on postoperative day 4 compared with the VEGF-treated postoperative day 10 group (P < 0.001) and the untreated postoperative day 4 group (P = 0.005). Qualitative analysis showed positive TUNEL signal in both pneumocytes and endothelial cells in both of these groups (Figs. 5 and 6). There were minimal TUNEL-positive cells in the untreated postoperative day 4 group and the VEGF-treated postoperative day 10 group. All slides were analyzed using confocal microscopy.


Figure 5
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  		Fig. 5. Median counts of terminal deoxynucleotidyl transferase biotin-dUTP nick end labeling (TUNEL)-positive cells per high-powered field (x63) on postoperative days 4 and 10 in mice undergoing partial PTX ± VEGF, with interquartile range in parentheses. There was no apoptosis in sham-operated mice.

 

Figure 6
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  		Fig. 6. Lung sections were stained as described for nuclei (blue), TUNEL (green, arrows), and CD31 (red). Postpneumonectomy lung sections are displayed ± VEGF. TUNEL-positive cells were widespread on postoperative day 10 without VEGF and postoperative day 4 with VEGF, correlating with the point of maximal compensatory lung growth in the growth curve.

 
Qualitative analysis of proSPC staining of type II pneumocytes under confocal microscopy showed that TUNEL stain colocalization involved but was not limited to the proSPC-positive cells (data not shown).

Increased proliferation was observed at timepoints leading up to complete compensatory growth in both the control and VEGF-treated groups. The most prominent staining was observed in the cells lining the small to medium-sized bronchioles (data not shown).

Vascular density. There were no differences in arterial density between the postop day 4 pneumonectomy +VEGF and the postop day 10 pneumonectomy groups (P = 0.7; data not shown).


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Liver parenchymal regeneration is characterized by angiogenic regulation, with endothelial cell proliferation promoting growth and apoptosis stopping it. Our laboratory showed that in the liver model, angiogenic factors such as bFGF and TNP-470 can regulate endothelial cell proliferation and apoptosis, and control growth (10). Given the vascular milieu of the lung, a similar angiogenesis-dependent control mechanism during compensatory lung growth might be expected. Compensatory lung growth in the mouse is characterized by a steady period of tissue growth, which is complete at 10 days. At this time there occurs a wave of both endothelial cell and pneumocyte apoptosis (Fig. 6). VEGF truncated this time course to 4 days with the same termination event (i.e., apoptotic wave). The exact temporal relationship between these two cell populations regarding apoptosis is not yet known.

We propose that exogenous VEGF stimulates increased blood vessel growth, thereby increasing energy substrate availability to lung tissue and hastening compensatory growth of the right lung. As noted, VEGF accelerated compensatory growth but did not cause overgrowth of the lung. In addition, morphometric studies, which showed no significant difference in air space volume between groups, suggest that tissue growth is mediated by hyperplasia, as opposed to hypertrophy, i.e., the formation of new cells rather than tissue "swelling." Further supporting that tissue growth was enabled by the creation of new cells; proliferation was increased in both control and VEGF-treated groups at the timepoints leading up to complete compensatory lung growth. It appears that apoptosis in our model is a more global pulmonary process as opposed to a cell-specific phenomenon, indicated by TUNEL positivity but not exclusivity in type II pneumocytes. Vascular development was also similar in VEGF-treated and normal compensatory growth cohorts, as indicated by arterial density measurements. Contrary to the studies in liver regeneration, bFGF had no effect on compensatory lung growth; compensatory growth was neither accelerated nor did it stimulate growth beyond its expected size. This suggests that the profiles of angiogenic factors controlling growth are distinct between these organ systems. It also provides a selection criterion that may prove to have therapeutic benefit in the future.

There was no effect seen with the two selective VEGF receptor inhibitors, MF-1 (anti-VEGFR-1) and DC101 (anti-VEGFR-2), suggesting that VEGF is not the sole regulator of compensatory lung growth. An alternative explanation may be that there was inadequate bioavailability of drug. The former appears more likely in that other agents, such as insulin-like growth factor, have been shown to be important in lung development and possibly compensatory lung growth (22, 24). Furthermore, treatment with small molecule chemical inhibitors of the two VEGF receptors failed to have an effect on compensatory lung growth, suggesting that the VEGF receptor antibodies were reaching their targets and confirming that blockage of these receptors is not sufficient to inhibit growth.

It is also worth noting that the same or lower doses of the drugs have proven to be effective at inhibiting angiogenesis during tumor growth, inflammation, and other VEGF-dependent physiological processes (1, 23, 35). Indeed, these data along with the ineffectiveness of bFGF to accelerate growth are important in characterizing the factors specifically involved in compensatory lung growth. It has been shown that vascular endothelium is heterogeneous with specific receptors expressed differentially depending on tissue type as well as physiological state (i.e., growth) (27, 28, 37). These phenotypic differences may account for the selective modulation of lung regeneration by VEGF and apparent ineffectiveness of bFGF and the various VEGF receptor inhibitors. As mentioned before, by delineating the unique traits of this system, directed therapeutic options can be designed.

It is important to note that although the objective evidence supports that VEGF is capable of accelerating compensatory lung growth, its role in normal physiological processes is largely unknown. Indeed, the fact that our various VEGF receptor inhibitors had no effect on the growth curves suggests that this role could be minimal.

Comparing the liver and lung organ systems, hepatic growth stops with a wave of endothelial cell death and hepatic parenchymal quiescence, whereas pulmonary growth stops with a wave of both endothelial cell death and pneumocyte death. In our model, the relationship between vasculature and its associated tissue is unknown. There are three possibilities: 1) endothelial cell apoptosis triggers pneumocyte apoptosis, 2) pneumocyte apoptosis reduces tissue mass and triggers a regression of excess endothelial cells, or 3) endothelial cell apoptosis and pneumocyte apoptosis occur concurrently. Our ability to manipulate the process of compensatory growth with VEGF suggests that endothelial cell regulation is the critical factor. The precise signaling mechanisms and temporal relationship need to be further elucidated.

Our studies suggest that angiogenesis plays an important role in compensatory lung growth. Exogenously administered VEGF accelerates growth. Similar to liver regeneration, angiogenesis modulators may control compensatory lung growth via the regulation of endothelial cell proliferation and apoptosis. However, exogenous VEGF has also been noted to have a stimulatory effect on lung epithelial cells (2), so it is possible that the enhancement effect we observed is multifactorial. As noted previously, type-2 pneumocytes and epithelial cells express VEGF receptors (19), which may function in a feedback mechanism or positive signaling for growth. HIF-2{alpha} has also been shown to be important in the appropriate vascularization of alveolar septa and vascular remodeling, as well as being expressed highly in alveolar pneumocytes during critical points in pulmonary maturation (5). It is likely that the VEGF mechanism of enhanced lung regeneration that we observed is the result of an effect on both angiogenesis and pneumocytes, amplifying the physiological process seen in normal lung development that is mediated by endogenous VEGF production by alveolar cells. This is further evidenced by the fact that we see apoptosis in both cell populations at the cessation of the regeneration period in both untreated and VEGF-treated cohorts.

The ability to manipulate this process with angiogenic agents may translate into therapies for pulmonary diseases which are related to lung immaturity, such as pulmonary hypoplasia secondary to congenital diaphragmatic hernia and broncho-pulmonary dysplasia. Typically, infants so afflicted die soon after birth. Those who do not succumb to their illness remain on prolonged mechanical ventilation or extra-corporeal membrane oxygenation, which themselves cause further pulmonary injury. Proangiogenic factors may be utilized to accelerate prenatal or postnatal lung growth and hence, prevent the morbidity and mortality of these treatment modalities. In addition, this study may provide insights into the mechanism underlying the development of pulmonary cavitary lesions and life-threatening hemoptysis, a potentially fatal complication observed in trials of anti-angiogenic agents for lung cancer (16). Studying the interplay between vasculature and tissue growth may give us unique insights into neoplastic derangement. Finally, the observation that lung and hepatic regeneration are controlled, at least in part, by different angiogenic factors highlights the importance of investigating this mechanism in different organ systems and raises the possibility that therapeutic approaches can be developed for stimulating regeneration in an organ-specific manner.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by the Howard Hughes Medical Institute SPORE Pilot Research Project and the Children's Hospital Boston Surgical Foundation.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. Puder, Children's Hospital Boston, 300 Longwood Ave., Boston, MA 02115 (e-mail: mark.puder{at}childrens.harvard.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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