1Respiratory Disease Area, Novartis Institutes for BioMedical Research, Horsham RH12 5AB; 2Molecular Pharmacology Group, Division of Biochemistry & Molecular Biology, Institute of Biomedical & Life Sciences and 4Department of Medicine, University of Glasgow, Glasgow G12 8QQ, Scotland, United Kingdom; 3Informatics and Knowledge Management, Novartis Institutes for BioMedical Research, CH-4002 Basel, Switzerland; and 5The Unit for Lung Investigations, Institute of Experimental and Clinical Medicine, Tallinn, 13419 Estonia
Submitted 10 November 2003 ; accepted in final form 16 March 2004
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ABSTRACT |
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cAMP-specific phosphodiesterase family type 4; chronic obstructive pulmonary disease; inflammation
There is a paucity of effective therapies for COPD (10). In contrast to asthma, the inflammatory response in COPD is largely insensitive to corticosteroid treatment (1). There has been considerable interest in cAMP-specific phosphodiesterase family type 4 (PDE4) as a therapeutic target for COPD (5, 10, 18, 35). Promising clinical data have been generated using both Cilomilast (Ariflo) (8, 40) and Roflumilast (BY-217) (10). The development of PDE4 inhibitors has been based on a large body of evidence from many laboratories that show such compounds can exert potent anti-inflammatory actions on a wide range of cells in vitro and in a range of animal models in vivo (4, 13, 39). However, one factor constraining the therapeutic use of many PDE4 inhibitors is their ability to cause nausea, which has placed limitations on the dosing levels that can be employed and hence their anti-inflammatory efficacy (5, 10, 18, 35). Furthermore, previous reports demonstrated how nonselective PDE4 inhibition, using the prototypic inhibitor rolipram, has the potential to lead to a diminished host defense response in animal models (2).
Currently, 11 different PDE families have been described, representing more than 50 different cAMP and cGMP PDE variants (36). PDE4 activity alone is provided by a large family of isoforms that are encoded by four genes (PDE4A, -4B, -4C, and -4D) (6, 15, 16). Each isoform is characterized by a unique NH2-terminal region, which is often involved in their intracellular targeting and may also serve to regulate their catalytic activity (16). These isoforms fall into three separate categories: the so-called "long" isoforms, which have the upstream conserved region (UCR)1 and UCR2 regulatory modules linking the isoform-specific NH2-terminal regions to the catalytic unit; the "short" isoforms, which lack UCR1; and the "supershort" isoforms that both lack UCR1 and have a truncated UCR2. These UCR modules can interact with each other and also serve to influence the functioning of the PDE4 catalytic unit where they orchestrate the functional consequence of phosphorylation of the PDE4 catalytic unit by extracellular signal-regulated kinase and of phosphorylation of UCR1 by cAMP-dependent protein kinase (PKA) (7, 15, 16). These differentially targeted and regulated isoforms are expressed in a cell type-specific fashion (15, 16), implying that individual PDE4 isoforms play specific functional roles. Indeed, differences in the functional role of isoforms have been implied by antisense strategies (28) and by selective inhibition (27, 29), and, additionally, gene knockout studies have shown that the PDE4B and PDE4D gene families can perform specific roles (12, 21).
Tenor et al. (37) have described cAMP PDE enzymatic activities in human alveolar macrophages from normal and atopic asthmatic donors and concluded that there was no difference between disease states. The expression of PDE4A, PDE4B2, PDE4C, and PDE4D has previously been analyzed for peripheral blood leukocytes of normal and atopic subjects (9), whereas more recently, the identification of PDE4A, PDE4B, and PDE4D subtypes in CD4+ and CD8+ lymphocytes from healthy and asthmatic subjects has been noted (23).
To date, no studies have been reported describing the distribution of PDE4 isoforms in inflammatory cells of patients with COPD. An ability to define which PDE4 isoforms and splice variants are expressed in inflammatory cells and whether there is a differential regulation in COPD vs. non-COPD as well as in smokers vs. nonsmokers is of great interest as it may contribute to our understanding of the basic molecular pathology of this respiratory disease. Such analyses may also pinpoint particular PDE4 variants as possible new therapeutic targets with less potential for dose-limiting side effects or potential host defense impairment.
This study was designed to investigate in detail the expression profile of 12 known PDE4 subtypes and splice variants as well as of PDE7A, PDE3A, and PDE3B subtypes in inflammatory cell types of relevance to COPD. In particular, the expression profile was analyzed and compared for lung macrophages, peripheral blood monocytes, T lymphocytes, and neutrophils of smokers with and without COPD, as well as for peripheral blood monocytes, T lymphocytes, and neutrophils of nonsmokers.
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MATERIALS AND METHODS |
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Collection of clinical samples. Clinical samples were obtained from volunteer subjects who had given their informed and written consent. Approval for the study was given by the local ethics committees (Guys Drug Research Unit and the Institute of Experimental and Clinical Medicine). The study conformed to the declaration of Helsinki.
Peripheral blood and bronchoalveolar lavage (BAL) samples were obtained from two groups (groups 1 and 2) of subjects at either the Guy's Drug Research Unit (London, UK) or the Institute of Experimental and Clinical Medicine (Tallinn, Estonia). Additional blood samples were taken from five nonsmoking subjects (group 3) from a volunteer donor panel at the Novartis Horsham Research Center. Group 1 consisted of 11 smokers diagnosed with mild to moderate COPD according to the Global Initiative for Chronic Obstructive Lung Disease criteria (11a). Group 2 was 10 smokers matched to the subjects in group 1 but diagnosed without COPD, as judged by a FEV1/FVC (forced expiratory volume in 1 s/forced vital capacity) ratio of >70%. The inclusion criteria for group 1 subjects were as follows: male or female COPD patients ages 4560 years; nonatopic; current smokers (>5 pack-years, having smoked for >10 years and currently smoking >5 cigarettes per day); airflow limitation (FEV1 6070% predicted for age and height, FEV1/ FVC ratio <70%) and with <10% reversibility to salbutamol; able to withhold bronchodilator therapy for at least 48 h; if female, using a medically acceptable form of contraception and having a negative result for pregnancy test before bronchoscopy, or be surgically sterile; and willing and able to give written informed consent.
The inclusion criteria for group 2 subjects were the same as for group 1 subjects, but they were diagnosed as not having any airflow limitation (FEV1/FVC >70% predicted for age and height) and with <10% reversibility to salbutamol.
The exclusion criteria for both groups were as follows: having taken regular inhaled or topical nasal corticosteroids in the preceding 3 mo; having received a course of prednisolone in the preceding 3 mo; having taken theophyllines within 14 days and other bronchodilators with 48 h of the scheduled bronchoscopy; having experienced an upper or lower respiratory tract infection within the preceding 4 wk; having chest X-ray appearances in previous month suggesting bilateral neoplasia, consolidation, or collapse; having an alcohol intake >28 units per week; participation in a clinical study within the last 3 mo; morbid obesity; and any clinically relevant history of serious cardiac or immunological disorder or other respiratory diseases as determined by the investigator. Table 1 summarizes the relevant clinical data for all subjects sampled for this study. Each subject from groups 1, 2, and 3 donated 100 ml of peripheral blood from which monocytes, T cells, and neutrophils were isolated. Subjects from groups 1 and 2 underwent a bronchoscopy with 150 ml of saline to provide BAL fluid (BALF) from which macrophages were isolated. Bronchoscopy was performed according to international guidelines (42), and BAL was obtained from the right middle lobe using three 50-ml aliquots of warmed saline. Patients fasted for 8 h before bronchoscopy. The procedure was performed under light sedation with 2.510 mg iv of midazolam together with 0.30.6 mg iv of atropine. Upper airway lignocaine topical anesthesia was applied. The dose did not exceed 6 mg/kg of lignocaine to avoid lignocaine toxicity from systemic absorption. Continuous administration of 2 l/min of supplemental oxygen was maintained throughout the procedure and for 60 min afterward. Nebulized salbutamol was administered as required after the procedure was completed to relieve bronchospasm. Lung function measurement was repeated once the patient had recovered from sedation and before discharge on the following day.
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Isolation of monocytes, T cells, and neutrophils from peripheral blood. Blood samples were collected using either 50-ml Falcon tubes containing 2 ml of filter-sterile 3.8% wt/vol trisodium citrate or using 10-ml heparin vacutainer tubes. Peripheral blood mononuclear cells (PBMCs) were separated from granulocytes and red blood cells using a Ficoll gradient essentially according to the manufacturer's instructions. Briefly, the blood was diluted 1:1 with Ca2+/Mg2+-free PBS. Up to 25 ml of this mixture was layered carefully onto 15 ml of Ficoll-Paque in a 50-ml Falcon tube at room temperature. A Ficoll gradient was established by centrifugation at 400 g for 30 min at room temperature. The layer containing PBMC was recovered and transferred to fresh ice-cold Falcon tubes. The remaining serum and Ficoll was aspirated to leave the red cell pellet. T cells and monocytes were purified sequentially from the PBMC layer, whereas neutrophils were purified from the red cell pellet, as described below. Blood cells in the recovered PBMC layer were washed twice with ice-cold MACS buffer (PBS + 0.4% bovine serum albumin + 2 mM EDTA), counted, and resuspended in 80 µl of MACS buffer for every 107 cells present. To this, 20 µl of each CD4 and CD8 multisort MACS beads were added, and T cells were separated using a MACS VS+ column as per the manufacturer's instructions. Monocytes were purified from the first flow-through fraction of this procedure using CD14 MACS microbeads. Neutrophils were isolated from the red blood cell pellet generated by the Ficoll gradient. The cell pellet was drained, and red blood cells were selectively lysed twice by resuspension in cold sterile H2O for exactly 30 s, immediately followed by the addition of an equal volume of 2x PBS, before centrifugation. The remaining white blood cells were washed in 20 ml of MACS buffer and resuspended in a final volume of 50 µl of MACS buffer for every 5 x 107 cells present. Neutrophils were recovered by adding an equal volume of CD16 MACS microbeads followed by separation on a type C MACS column as described by the manufacturer. In all cases, the recovered cells were counted and centrifuged at 400 g for 5 min at 4°C, and the supernatant fraction was discarded. The recovered pellets were lysed in 60 µl of RLT buffer for every 106 cells present, but using not less than 350 µl. Samples were kept chilled at all times to minimize degradation. The lysate was thoroughly disrupted by pipetting up and down and flash-frozen at 70°C.
Extraction of total RNA and cDNA synthesis.
For all cell types isolated, frozen lysates were warmed at 37°C for 10 min and homogenized using QIAshredder columns (Qiagen) according to the manufacturer's instructions. RNA was extracted from the homogenized lysates using QIAgen RNeasy mini or midi kits following the kit instructions. The RNA was treated with RQ1 DNase (1 U/µl) to remove genomic DNA contamination. RNA (75 µl) was mixed with 20 µl of RQ1 and 5 µl of 20x RQ1 buffer (0.6 mM Tris·HCl, 72 mM MgCl2, 120 mM NaCl) and incubated at 37°C for 3045 min. After this, the RNA was repurified using the "clean-up" protocol of the RNeasy mini kit. To check for successful genomic DNA removal, each sample was tested in RT-PCR for -actin with and without reverse transcriptase using Promega Access RT-PCR kit.
Once samples were confirmed as DNA free, first-strand cDNA was synthesized using the Pharmacia first-strand cDNA synthesis kit. Five micrograms of RNA were diluted to 20 µl using nuclease-free water and then heated to 65°C for 10 min. To this, 14 µl of a mix containing random primers, oligo(dT), buffer, and enzyme (supplied with kit) were added. This was incubated for 1 h at 37°C followed by a 90°C incubation for 5 min to inactivate the reverse transcriptase. The completed reaction was diluted with nuclease-free water to 1 ml to allow addition of 5 µl (equivalent to the addition of 25 ng of RNA) for subsequent PCR.
Specific amplification by PCR and primer design.
First-strand cDNA was used as a template for PCR analysis of the following PDE isoforms and splice variants: PDE4A (generic) and splice variants PDE4A4, PDE4A10, and PDE4A7 (2EL); PDE4B (generic) and splice variants PDE4B1, PDE4B2, and PDE4B3; PDE4C (generic); PDE4D (generic) and splice variants PDE4D1, PDE4D2, PDE4D3, PDE4D4, and PDE4D5; PDE3A (generic); PDE3B (generic); and PDE7A (generic). PCR primers were designed to cover either a conserved region within a PDE isoform (generic) or the unique 5' end of the mRNA to target a specific splice variant. The sequences used were obtained from the GenBank database, and GenBank accession numbers are shown in Table 2. Additional primers were designed to amplify -actin and transferrin as amplification controls and for normalization purposes. Primers were designed using the Applied Biosystems software Primer Express and selected for their position within the sequence and calculated annealing temperature. Amplified PCR fragments were fractionated and visualized on a 2% agarose gel in 1x TAE buffer (40 mM Tris-acetate and 1 mM EDTA) containing 1 µg/ml of ethidium bromide. Sequences of PCR primers used are listed in Table 1. Amplification conditions and primer sequences were tested and optimized on first-strand cDNA synthesized from total brain RNA (Clontech). Reaction conditions were optimized to enable detection in as little as 25 ng of total RNA. Brain RNA similarly converted to cDNA served as positive control for all PCR reactions as all tested PDE variants were found to be present in brain tissue. PCR reactions were performed on each sample in duplicate or triplicate using the following conditions: 18.75 pmol of each forward and reverse primer; 12.5 µl of 2x PCR mix; 5 µl of the cDNA synthesis reaction equivalent to 25 ng of RNA; and nuclease-free water to a total volume of 25 µl. Amplifications were performed using either Epicentre 2x PCR mix H and 0.625 units of Epicentre MasterAmp Amplitherm DNA polymerase for amplification of PDE4A4, PDE4B3, and
-actin or Qiagen HotStarTaq Mastermix for all other reactions and subjected to the following program on a Biometra T3 Thermocycler: 95°C for 5 min; 40 cycles of 94°C for 30 s, 60°C for 30 s, and 72°C for 1 min; a final extension of 72°C for 10 min; followed by pause at 4°C. Semiquantitative RT-PCR was performed in duplicate by halting the PCR amplification during the exponential phase as determined for each variant in preliminary experiments. Preliminary experiments had shown that
-actin was a better and more constant standard for comparisons. PCR products were separated on a 2% agarose gel. DNA bands were visualized by ethidium bromide staining, and their intensities were normalized vs.
-actin. Fluorescent intensities were measured and analyzed using ImageQuant software (Molecular Dynamics). Real-time quantitative PCR was performed using an ABI 7900 instrument. The PDE4A4 primer and probe sequences were as follows [forward: 5-CGCACCGGCCCATAGAG-3'; reverse: 5'-TGCCAGTGCCATGGAAGGA-3'; 6-carboxyfluorescein dye (FAM)-probe: 5'-ACCCGCATGTCCTG-3'] and were used at the concentrations of 800 and 250 nM, respectively. Control reagents for
-actin, including primers and VIC-reporter probe, were purchased from ABI. Primers were used at 900 nM and probe at 200 nM.
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Statistical analysis. Statistical analysis of the semiquantitative RT-PCR results was performed using Mann-Whitney's or Wilcoxon's test and Student's t-test or paired t-test as indicated.
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RESULTS |
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Three groups of subjects were recruited. Group 1 consisted of 11 smokers diagnosed with COPD, whereas group 2 consisted of 10 smokers matched for age, gender, and smoking history (see MATERIALS AND METHODS) but without COPD. Five samples in each group were used for RNA profiling work, and the remaining samples in each group were used for enzyme activity determinations. Group 3 was made up of five nonsmokers who volunteered to donate peripheral blood samples for PCR analysis. Because elevated cAMP levels have been shown to affect the expression of certain PDE4 variants (24, 30), care was taken to avoid exposure to cAMP-elevating agents (e.g., 2-adrenoceptor bronchodilators) before sampling.
Evaluating primer pairs. To undertake this study, gene-specific amplification conditions and primer pairs needed to be optimized and validated for each PDE4 splice variant. Human total brain RNA was used for this purpose and was included as a positive control for all the RT-PCR experiments. Table 2 shows the primer sequences used for RT-PCR and amplicon sizes, and Fig. 1 illustrates schematically where the PDE subtype generic and isoform-specific RT-PCR primers are located. Figure 2 shows a typical agarose gel with the RT-PCR expression profile for the 15 PDE variants in whole human brain. All amplicons migrated at the expected rate. As expected from the primer design, the PDE4D1/2 pattern produced two bands, as a fragment from both PDE4D1 and from PDE4D2 was amplified. All amplicons were further subjected to a diagnostic restriction enzyme analysis to confirm the identity of the fragments produced (data not shown). Bands on the agarose gel were scored as positive only when the amplification was reproducible in each PCR replicate. Splice variant-specific amplicons for PDE4 were scored as positive only when the expected fragment for the corresponding generic PDE4 subtype was also amplified in the same sample.
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Comparing PDE4 expression patterns among smokers with and without COPD by semiquantitative and quantitative RT-PCR. First, a semiquantitative RT-PCR approach was used to address putative disease-specific or cell type-specific differences observed in the qualitative RT-PCR analysis of PDE4 transcript levels.
When doing this, the transcripts of the PDE4A4 long isoform, when normalized to -actin control, were found to be significantly upregulated in BAL macrophages from COPD patients compared with non-COPD subjects (P < 0.01, Fig. 4). In contrast, PDE4A4 expression was not seemingly different in blood monocytes of the same COPD subjects compared with non-COPD controls (Fig. 4). Interestingly, this was the only significant difference among the 15 PDE variants analyzed in inflammatory cells of smokers with COPD vs. smokers without COPD. It was, therefore, decided to further confirm the differential expression of PDE4A4, specifically in COPD vs. healthy smokers, by real-time quantitative PCR. The data in Fig. 5 confirm a significant (P < 0.02) upregulated expression for PDE4A4 in BAL macrophages of smokers with COPD.
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DISCUSSION |
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Four genes encode at least 12 different PDE4 splice variants, and for those analyzed here, we show clear evidence for specific tissue distribution (6, 7, 15). Gene knockout studies (12, 21), selective inhibition (27, 29), antisense (28), and dominant negative strategies (33) imply that specific PDE4 isoforms may have particular functional roles. This is considered most likely achieved (7, 16) by their specific intracellular targeting, leading to the control of anchored PKA (36). A number of studies have been published describing the expression of particular PDE4 variants in leukocytes (9, 23). However, a detailed and comprehensive investigation of the expression and distribution of PDE4 splice variants in inflammatory cells of relevance to COPD has not been reported. In addition, no information is available on the PDE4 isoform expression profile in lung bronchoalveolar macrophages, and there is also no information on the comparison of expression in inflammatory cells obtained from smokers with COPD vs. smokers without COPD and vs. nonsmokers.
PDE4 enzymes have a very high specific activity and are expressed at very low levels endogenously (4, 7, 16). This makes it extremely challenging to detect native levels of expression, especially under conditions where cell numbers and samples are very limited, as in clinical studies. Furthermore, there is a paucity of effective isoform-specific antisera, and certain major isoforms have proved to be remarkably difficult to generate antisera against (see e.g., Ref. 15). RT-PCR provides an extremely powerful and sensitive means of assessing transcripts for proteins. Using qualitative RT-PCR analysis, we show that all cell types tested here have transcripts for at least one PDE4 variant representing each PDE4 subfamily. The only exception was T cells, which did not express any PDE4C, as measured by RT-PCR. In many cases, several splice variants representing a PDE4 subtype were present. Furthermore, none of the cells studied here expressed just a single PDE4 subtype. Additionally, we noted that PDE4D5 was absent in BAL macrophages of both COPD and non-COPD subjects, whereas this long isoform was clearly present in the monocytes from the same individuals. PDE4D3 was not observed in blood neutrophils but was detected in all other cell types.
Semiquantitative analyses revealed disease and cell type-specific expression patterns for certain PDE4s. Thus PDE4A4 was specifically and uniquely upregulated in lung macrophages of COPD subjects, whereas PDE4A4 and PDE4B2 were both upregulated in peripheral blood monocytes from smokers compared with nonsmokers (Figs. 4C and 6). PDE4D5 was downregulated in lung macrophages compared with blood monocytes from the same subjects, irrespective of the disease state (Fig. 8). Furthermore, PDE4C was very poorly expressed in blood monocytes, yet clearly upregulated in the macrophages of the same individuals, again irrespective of disease state. Real-time quantitative PCR further confirmed the upregulation of PDE4A4 in COPD subjects vs. healthy controls (Fig. 5).
Previous RT-PCR data (41) have indicated that PDE4B2 transcripts predominate in peripheral blood monocytes and neutrophils from healthy volunteers. Indeed, it was suggested that PDE4B2 constituted 80.2% of the PDE4 population in unstimulated monocytes and 99.7% in neutrophils, with PDE4D forming 0.3%, and PDE4A and PDE4C together <0.1% of the total PDE4 transcripts in neutrophils. Our data are clearly at variance with this, showing that transcripts for all four PDE4 subfamilies were readily and abundantly detected in both monocytes and neutrophils (Fig. 3). Moreover, transcripts for PDE4B1, in addition to PDE4B2, were clearly expressed in both monocytes and neutrophils as well as in T cells and BAL macrophages (Fig. 3). Transcripts for multiple splice variants within the PDE4A and PDE4D subfamilies were also clearly evident. The reasons for these differences are not clear but could include the use of suboptimal PCR conditions or primer sequences, as we found it was essential to thoroughly validate the primers and conditions used not only against plasmids encoding specific isoforms but especially on whole brain RNA, which we show here provides an excellent source of the full range of known PDE4 isoforms (Fig. 2). Furthermore, whereas LPS activation of monocytes has been shown to increase PDE4B2 transcript levels (26), we show here that, using peripheral blood monocytes and lung macrophages obtained from smokers with and without COPD, a difference in expression levels for PDE4B2 could not be observed between disease states and cell types using semiquantitative RT-PCR. This may indicate a difference from in vivo activation compared with that seen in vitro with LPS.
PDE4C is often ignored as contributing to inflammatory cell function based on its low-level expression in these cells. We confirm that PDE4C transcript levels are lower compared with those of other PDE4 subfamilies in blood monocytes and neutrophils and confirm that PDE4C transcripts are not found in blood T cells (Fig. 3) (11). However, PDE4C mRNA is clearly present (Fig. 6) in elevated amounts in BAL macrophages compared with monocytes from the same individuals. Although no differential expression was observed between COPD and healthy subjects, PDE4C cannot be excluded as a possible macrophage therapeutic target for COPD.
Transcripts for PDE3A, PDE3B, and PDE7A were detected in all four cell types analyzed, extending observations of their expression in primary CD4+ and CD8+ T lymphocytes (11). PDE3A was very weakly present, especially in lung macrophages and blood monocytes. PDE4A7 is a curious isoform that is NH2- and COOH-terminally truncated to such an extent that it is catalytically inactive (14). We clearly identified transcripts for it in a wide variety of blood cells, suggesting that further investigation of this isoform is warranted.
A key finding of this study is the specific upregulation of PDE4A4 in lung macrophages from COPD subjects. A key property of PDE4A4 is its ability to interact with the SH3 domains found in various Src family tyrosyl kinases involved in activation of immune system cells (32). This interaction serves to both target PDE4A4 to specific intracellular locations as well as affect the conformation of the catalytic unit indicated by a heightened sensitivity to inhibition by the archetypal PDE4-selective inhibitor rolipram (32). Importantly, the activity of PDE4A4 increases upon activation of U-937 monocytic cells by proinflammatory agents such as LPS and interferon- (27). This suggests a key role for this isoform in lowering localized cAMP levels to facilitate inflammatory cell activation. Additionally, overexpression of PDE4A5, the rodent homolog of PDE4A5, has been shown to protect fibroblasts against apoptosis (19), which might again suggest that inhibition of this isoform is of potential therapeutic importance in attenuating inflammatory lung disease. Intriguingly, PDE4A5 has also very recently been shown to interact specifically with the immunophilin XAP2 (3). Although the functional consequences of this are still unknown, we note that XAP2 is also able to interact with the aryl hydrocarbon (dioxin) receptor, which can be activated by pollutants (22).
Given the significant upregulation of PDE4A4 in BAL macrophages of smokers with COPD, we also measured the PDE4 enzyme activity in BAL macrophages of smokers with and without COPD. The data show that the total PDE4 activity is clearly upregulated in BAL macrophages of smokers with COPD. Unfortunately, we were unable to measure activity specifically associated with PDE4A. This may be due to low absolute amounts of PDE4A protein or due to poor immunoprecipitating capacity of the antiserum used. Although transcripts for PDE4A4 were markedly upregulated in these samples and this possibly contributed to the increased enzyme activity, we cannot rule out contributions from other PDE4 variants and/or increases due to posttranslational activation of all or some of the PDE4 variants. Nonetheless, the observation that the total PDE4 enzyme activity is markedly increased in macrophages of COPD subjects compared with non-COPD subjects is of interest.
Because macrophages are a hallmark of inflammation in COPD (1, 20), the finding of a highly specific and significant differential expression of PDE4A4 in macrophages of COPD subjects may well be of importance. Indeed, the results of this study lead us to suggest that PDE4A4 may be a PDE4 isoform-specific therapeutic target for COPD. Whereas PDE4A4 is clearly also expressed in other cell types, including immune cells, its disease-specific upregulation in lung macrophages could possibly provide a new route into a PDE4-targeted anti-inflammatory treatment for COPD, with less potential for dose-limiting side effects. Testing the new hypothesis that the PDE4A4 isoform is a plausible new therapeutic target for COPD will require a new emphasis on either making specific inhibitors for PDE4A4 or devising new strategies to disrupt PDE4A4-specific intracellular interactions (19, 27, 32) to help elucidate its role in COPD.
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FOOTNOTES |
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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.
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REFERENCES |
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