Acetazolamide Serves as Selective Delivery Vehicle for Dipeptide-Linked Drugs to Renal Cell Carcinoma

The majority of cytotoxic agents used for the pharmacotherapy of cancer do not preferentially accumulate at the tumor site, leading to potential toxicities and to suboptimal therapeutic efficacy (1–4). In tumor-bearing mice, unfavorable tumor:organ ratios have been reported for many drugs, including doxorubicin (5), paclitaxel (6), cisplatin (7), cyclophosphamide (8), sunitinib (9), and different fluorinated pyrimidines (10), to name just a few. For example, the relative uptake of doxorubicin in the tumor (expressed as percentage injected dose per gram or %ID/g) was found to be only 5% to 10% of the corresponding values in lung, heart, or liver, in the time period 0.5 to 8 hours (5). Importantly, similar trends are emerging from the positron emission tomography study of patients with cancer, who had received radiolabeled preparations of cytotoxic drugs (3, 4).

The coupling of potent cytotoxic payloads to antibodies or small organic molecules, acting as selective pharmacodelivery vehicles, has been proposed as a general strategy to improve the therapeutic index of anticancer drugs. Indeed, certain antibody–drug conjugates (ADCs; refs. 11–14) and small molecule–drug conjugates (SMDCs; ref. 2) have exhibited promising activity in preclinical models of cancer. Recently, 2 ADC products (Adcetris and Kadcyla) have gained marketing authorization for oncological applications.

Some ADCs cured tumors in mice at doses of 0.5 to 3 mg/kg, whereas the same products could be administered at 60-fold greater dose with acceptable safety profiles (15). Unfortunately, such a high therapeutic index is rarely observed in the clinic, suggesting that the tumor-targeting properties of the antibody, the drug release process, and/or the intrinsic sensitivity of tumor cells may be dramatically different in preclinical models and in patients with cancer. Nuclear medicine studies with radiolabeled antibody preparations have previously shown that lower tumor:organ ratios are often observed in humans compared with tumor-bearing mice, possibly reflecting differences in antigen abundance and vascular permeability (16, 17). SMDCs may represent an attractive alternative to ADC products, as smaller pharmaceuticals extravasate more rapidly and diffuse more homogenously within the neoplastic mass (2, 18–22). Unlike antibodies (which can easily be raised against the majority of target proteins of interest), it is not always easy to generate high-affinity small-molecule ligands to tumor-associated antigens. However, excellent tumor-targeting results have been reported for folate analogues targeting folate receptor–positive tumors (23, 24), substituted urea derivatives targeting prostate-specific membrane antigen (25), somatostatin antagonists targeting the somatostatin receptor (26) and for carbonic anhydrase IX (CAIX) ligands (16–18, 27).

It has generally been assumed that ADC and SMDC products would crucially rely on the use of ligands, capable of selective internalization of the conjugate into the tumor cells, followed by an intracellular liberation of the cytotoxic payload (11–14). This concept, however, has recently been challenged, as potent and selective anticancer activity has been observed with ADCs and SMDCs specific to antigens, which do not internalize (19, 20, 28–31).

CAIX is a membrane protein, expressed in the majority of clear cell renal cell carcinomas (32–34). CAIX is virtually undetectable in most normal adult tissues, except for certain structures in the gastrointestinal tract (stomach, duodenum, gallbladder), which are strongly positive for the antigen. Anti-CAIX monoclonal antibodies efficiently localize to metastatic renal cell carcinoma lesions in patients but, surprisingly, not to CAIX-positive stomach, duodenum, and gallbladder (35), suggesting that differences in vascular permeability may account for the process.

To the best of our knowledge, CAIX is a tumor-associated target which does not internalize (19), even though at least one anti-CAIX ADC product has been studied in the clinic in patients with cancer (36). CAIX-positive tumors can be efficiently targeted using certain small aromatic sulfonamides (18–20, 33, 37), including acetazolamide.

We have previously reported that acetazolamide can be used to deliver disulfide-linked derivatives of DM1 (a potent cytotoxic maytansinoid) to CAIX-positive tumors, leading to a potent antitumor effect (19, 20). We hypothesized that the extracellular drug release process at the tumor site would be facilitated by the death of tumor cells and the subsequent release of glutathione into the tumor extracellular space, which can work as a reducing agent on disulfide bonds, favoring an amplified cascade of drug release and tumor cell death. Since disulfide bonds can be unstable in vivo and since most anticancer drugs (unlike DM1) do not contain thiol groups suitable for coupling purposes, it would be interesting to learn whether potent noninternalizing SMDCs can be generated with stable peptidic linkers and with other classes of drugs. In this article, we describe that potent anticancer SMDCs can be generated by coupling acetazolamide to highly cytotoxic drugs (e.g., MMAE and PNU-159682), using a linker containing a valine–citrulline moiety and a self-immolative spacer. These findings reinforce the concept that potent antitumoral activity can be generated by the efficient release of toxic payloads in the tumor extracellular space, without ligand internalization into the target cells of interest.

Detailed synthetic procedures and characterization of the presented compounds (i.e., HPLC purity of final products, MS and NMR data, SPR assays) are described in the Supplementary Information (Supplementary Figs. S1–S11), together with additional biological data (i.e., stability of 4a in the presence of Cathepsin B and CAIX, biodistribution in CAIX-negative lesions), and protocols (i.e., expression and purification of recombinant human CAIX; Supplementary Figs. S12–S15).

Peptidic precursors of compounds 1–5 (Fig. 1) were synthesized by solid phase peptide synthesis (SPPS), using Fmoc-protected amino acids. Chlorotrityl resin (500 mg), preloaded with Fmoc-Cys(Trt)-OH, was swollen with DMF (10 ml) for 15 minutes inside a syringe equipped with a filter pad. Fmoc deprotection was achieved by shaking the resin with 20% v/v piperidine in DMF (5 mL) for 10 minutes for 3 times. After deprotection, the resin was washed with DMF (4 × 10 mL). Fmoc-protected amino acids (3 eq) were activated with HATU (3 eq) and DIPEA (6 eq) in DMF (5 mL) for 15 minutes in ice bath. After this time, the solution was allowed to react with the syringe for 1 hour and discarded and the resin washed with DMF (4 × 10 mL). Coupling and deprotection steps were alternated, to obtain the desired peptide sequence.

Acetazolamide or free amide moieties were then introduced by performing a CuAAC “click” reaction (38) with the corresponding peptide derivatives (carrying an azide moiety) on solid phase, followed by side chain deprotection, cleavage from the resin, and RP-HPLC purification.

Radiolabeling procedures with technetium-99m were performed essentially as described (18). Briefly, compound 1 (60 nmol) in TBS pH 7.4 (50 μL) was mixed with SnCl₂ (Sigma Aldrich, 200 μg) and sodium glucoheptonate (TCI, 20 mg) in H₂O (150 μL). TBS at pH 7.4 (600 μL) was added and the resulting solution degassed for 5 minutes by bubbling with nitrogen gas. The eluate from a ^99mTc-generator (200 μL, ca. 200 MBq, Mallinckrodt) was added and the reaction mixture heated to 90°C for 20 minutes. After cooling to room temperature, an aliquot was analyzed by RP-HPLC (XTerra C18, 5% MeCN in 0.1% aq. TFA to 80% over 20 minutes on a Merck-Hitachi D-7000 HPLC system equipped with a Raytest Gabi Star radiodetector). Technetium-99m incorporations >95% were routinely achieved. The radioactive solution was then diluted to the desired concentrations with a molar excess of compound 1, dissolved in TBS, pH 7.4.

Peptide precursors of compounds 2–5 were dissolved in PBS (50 mmol/L phosphate, 100 mmol/L NaCl, pH 7.4) and a solution of maleimidocaproyl-linker-p-aminobenzylalcohol-drug in DMF was added at suitable molar ratios. The mixtures were stirred at room temperature until completion and the solvents were removed under vacuum. The conjugates were purified from the crude material via RP-HPLC and lyophilized to obtain the solid products. Purity was checked by UPLC on a Waters Acquity UPLC H-Class System using a ACQUITY UPLC BEH C18 Column, 130 Å, 1.7 μm, 2.1 mm × 50 mm column. Purities higher than 95% were routinely achieved. Details related to specific conjugates are reported in the Supplementary Information.

The human renal cell carcinoma cell line SKRC-52 was kindly provided by Professor E. Oosterwijk in 2008 (Radbound University Nijmegen Medical Centre, Nijmegen, The Netherlands) and subsequently stored in liquid nitrogen. Upon thawing, cells were kept in culture for no longer than 14 passages, tested for CAIX expression by immunofluorescence, and not further authenticated. SKRC-52 and HEK293 (ATCC, CRL-1573) were maintained in RPMI medium (Invitrogen) supplemented with 10%, FCS (Invitrogen) and antibiotic/antimycotic (1%, AA, Invitrogen) at 37°C and 5% CO₂. For passaging, cells were detached using Trypsin-EDTA 0.05% (Invitrogen) when reaching 90% confluence and reseeded at a dilution of 1:6.

SKRC52 or HEK 293 cells were seeded into 4-well coverslip chamber plates (Sarstedt) at a density of 10⁴ cells per well in RPMI medium (1 mL, Invitrogen) supplemented with 10% FCS, AA, and HEPES (10 mmol/L) and allowed to grow for 24 hours under standard culture conditions. The medium was replaced with medium containing 2a or 2b (120 nmol/L), after 30 minutes or 1 hour, Hoechst 33342 nuclear dye (Invitrogen) was added and randomly selected cells imaged on an SP8 confocal microscope equipped with an AOBS device (Leica Microsystems).

Compounds 4a and 5a were dissolved in PBS (100 μg/mL) and incubated at 37°C in a shaking incubator. Aliquots (150 μL) were taken at different time points (0 minutes, 15 minutes, 1, 6, 24, 48 hours) and frozen at −20°C. Standard solutions of the conjugates were prepared in PBS at different concentration (25, 50, 100, 150 μg/mL) and stored at −20°C. The standards and the samples were analyzed in triplicate by liquid chromatography/mass spectrometry (LC-MS) on a Waters Xevo G2-XS QTOF coupled to a Waters Acquity UPLC H-Class System using a ACQUITY UPLC BEH C18 Column, 130 Å, 1.7 μm, 2.1 mm × 50 mm column. Peaks relative to the intact SMDCs were integrated, and area values from the standards were used to obtain calibration curves. Concentration of intact test compounds in the samples was determined at the different time points to obtain a stability profile of the conjugates.

To measure mouse serum stabilities of the conjugates, the compounds were dissolved in freshly thawed mouse serum (Invitrogen; 100 μg/mL) and incubated at 37°C in a shaking incubator. Aliquots of 150 μL were taken at different time points (0, 20 minutes, 1, 3, 6, 24 hours) and frozen at −20°C. Standard solutions were prepared in mouse serum (Invitrogen) at different concentration (25, 50, 100, 150 μg/mL) and stored at −20°C. All the samples and the standards were thawed and diluted with 4 volumes of MeOH. After vigorous vortex agitation for 1 minute, the protein precipitate was spun down and 400 μL of the supernatant were lyophilized overnight. The resulting solid material was re-dissolved in 100 μL of Millipore water and analyzed as described above.

SKRC-52 cells were seeded in 96-well plates in RPMI added with 10% FCS (100 μL) at a density of 5,000 cells per well and allowed to grow for 24 hours. The medium was replaced with medium containing different concentrations of test substance (1:3 dilution steps) and plates were incubated under standard culture conditions. After 72 hours, the medium was removed, MTS cell viability dye (20 μL, Promega) in 150 μL of the medium was added, the plates were incubated for 2 hours under culture conditions, and the absorbance at 490 nm measured on a Spectra Max Paradigm multimode plate reader (Molecular Devices). Experiments were performed in triplicate and average cell viability calculated as measured background corrected absorbance divided by the absorbance of untreated control wells. IC₅₀ values were determined by fitting data to the 4-parameter logistic equation, using a Prism 6 software (GraphPad Software) for data analysis.

All animal experiments were conducted in accordance with Swiss animal welfare laws and regulations under the license number 27/2015 granted by the Veterinäramt des Kantons Zürich.

SKRC-52 cells were grown to 80% confluence and detached with Trypsin-EDTA 0.05% (Life Technologies). Cells were washed with Hank balanced salt solution (HBSS, pH 7.4) once, counted and resuspended in HBSS to a final concentration of 3.4 × 10⁷ cells/mL. Aliquots of 5 × 10⁶ cells (150 μL of a suspension) were injected subcutaneously in the right flank of female athymic Balb/c nu/nu mice (8–10 weeks of age, Janvier).

SKRC-52 xenografted tumors were implanted into female Balb/c nu/nu mice (Janvier) as described above and allowed to grow for 3 weeks to an average volume of 0.25 mL. Mice were randomized (n = 5 per group) and injected intravenously with different doses of radiolabelled preparations (15–25 MBq, 70–560 nmol/kg). Mice were sacrificed 6 hours after the injection by CO₂ asphyxiation and organs extracted, weighted, and radioactivity measured with a Packard Cobra γ-counter. Values are expressed as %ID/g ± SD.

Female Balb/c nu/nu mice bearing subcutaneous SKRC-52 tumors were injected intravenously with Compound 3a, containing the near-infrared dye moiety IRDye680RD (LI-COR Biosciences; 250 nmol/kg), dissolved in sterile PBS (100 μL). Mice were anesthetized with isoflurane and fluorescence images acquired on an IVIS Spectrum imaging system (Xenogen, exposure 1 second, binning factor 8, excitation at 675 nm, emission filter at 720 nm, f number 2, field of view 13.1). Images were taken before the injection and after 10 minutes, 1, 3, and 6 hours. Food and water was given ad libitum during that period.

Recommended dose of compound 4a suitable for therapy experiments was determined by dose escalation in wild-type female athymic Balb/c nu/nu mice (8–10 weeks of age, Janvier). A schedule of 5 injections on 5 consecutive days was used to compare increasing doses (250 or 500 nmol/kg) of the targeted derivative 4a with untargeted compound 4b (Supplementary Fig. S13). Three mice were used for each group. Tolerated dose was defined when animals did not lose more than 5% of their initial body weight over the duration of the experiment after the initial injection.

SKRC-52 xenografted tumors were implanted into female Balb/c nu/nu mice (Janvier) as described above and allowed to grow for 2 weeks to an average volume of 0.1 mL. Mice were randomly assigned into therapy groups of 4 or 5 animals and treatment started by injecting a solution of the targeted drugs, untargeted drugs, or vehicle (PBS only or PBS containing 1% of DMSO) intravenously (tail vein) at the doses and with the schedules indicated in the text. Compounds 4a and 4b were injected as solutions in sterile PBS. Compounds 5a and 5b were injected as solutions in sterile PBS containing 1% DMSO. Animals were weighed and tumor sizes measured daily with an electronic caliper. The tumor volume was calculated according to the formula: (long side) × (short side) × (short side) × 0.5. Animals were sacrificed when the termination criteria were reached. Prism 6 software (GraphPad Software) was used for data analysis (regular 2-way ANOVA with the Bonferroni test).

SKRC-52 tumors were excised from the animals treated with conjugate 5a or alternatively with vehicle during the therapy experiment, embedded in OCT medium (Thermo Scientific), and cryostat sections (10 μm) were cut. Slides were stained using the following antibodies: mouse anti-human CAIX GT12 (Thermo Scientific) to detect the antigen and rat anti-mouse CD31 (BD Biosciences) to detect endothelial cells. Nuclear staining was performed with DAPI. Anti-mouse IgG-AlexaFluor488 (Molecular Probes by Life Technologies) and anti-rat IgG-AlexaFluor594 (Molecular Probes by Life Technologies) were then used as secondary antibodies for microscopic detection.

Acetazolamide was coupled to a Lys-Asp-Cys–based ^99mTc-chelating moiety, which had previously been used for the imaging of patients with cancer with folate derivatives (39), using a “click” chemistry reaction (Fig. 1). The resulting conjugate 1 could be labeled with ^99mTc (radiolabel incorporation >95% routinely achieved; 160–210 MBq/mL) for quantitative biodistribution analysis in tumor-bearing mice. Click chemistry was also used to install the acetazolamide moiety onto Asp-Arg-Asp-Cys peptidic derivatives, which were subsequently coupled to a red Alexa fluorophore (Compound 2a), a near-infrared dye (Compound 3a), or cytotoxic drugs, based on the monomethyl auristatin E (MMAE; Compound 4a) or PNU-159682 (Compound 5a) active moieties (Fig. 1). As negative controls devoid of CAIX binding, derivatives containing an amide function instead of the acetazolamide moiety were used (Compounds 2b, 4b, and 5b).

Confocal microscopy experiments were performed to investigate the internalization of our acetazolamide derivatives upon ligand binding to CAIX. SKRC-52 cancer cells (expressing high levels of CAIX) were incubated with compound 2a and the fluorescence derived from Alexa594-labeled acetazolamide was imaged after incubation for 30 and 60 minutes (Fig. 2). Compound 2a showed selective accumulation on the membrane SKRC-52 cells, as compared with CAIX-negative HEK 293 cells. No intracellular fluorescence was observed throughout the experiment, indicating that the AAZ binding to CAIX was not followed by a significant receptor-mediated endocytosis. In a control experiment, compound 2b, devoid of the targeting moiety, showed no interaction with CAIX-positive SKRC-52 cells (Fig. 2).

The tumor-targeting properties of acetazolamide derivatives were characterized using both radioactive and near-infrared fluorescence methods in nude mice bearing subcutaneously grafted SKRC-52 tumors (40). Compound 1, labeled with ^99mTc, was injected intravenously at doses ranging between 70 and 560 nmol/kg. A preferential tumor uptake was observed 6 hours after intravenous administration at all doses (Fig. 3A). However, a progressive decrease in percent injected dose per gram of tumor was detected, reflecting target saturation at the highest dose. Interestingly, compound uptake in normal organs (e.g., liver, lung, kidney, intestine, stomach) also decreased at higher doses. These biodistribution results suggested that doses around 200 to 300 nmol/kg may represent a good compromise between high tumor uptake and low accumulation in normal tissues. The near-infrared fluorescent derivative 3a, administered at 250 nmol/kg, was shown to preferentially accumulate in the tumor at various time points, between 5 minutes and 6 hours (Fig. 3B). These results are in keeping with the ones previously reported by our group with similar fluorescently labeled derivatives (17, 18, 30), indicating that the acetazolamide moiety can deliver different types of payloads to SKRC-52–positive tumors.

Prior to therapy studies, the stability of acetazolamide-drug conjugates 4a and 5a was investigated in vitro at 37°C, both in PBS and in mouse serum (Fig. 4). For both compounds, a half-life in PBS >48 hours was observed. A lower stability was seen in mouse serum, but >70% of both conjugates were intact after 6 hours (i.e., a sufficiently long time, compared with the circulatory half-life of acetazolamide derivatives in blood, which is typically shorter than 15 minutes; ref. 16). Mass spectrometric analysis of compound 4a revealed free MMAE as the main release product, whereas in the degradation profile of compound 5a in mouse serum, free PNU-159682 was not identified, probably due to low sensitivity of the mass spectrometer to a such hydrophobic moiety.

An in vitro toxicity study, performed with SKRC-52 cells incubated with the conjugates 4a,b-5a,b or with the corresponding unmodified drugs, confirmed that the acetazolamide coupling had led to the formation of prodrugs. Indeed, all the conjugates resulted to be less toxic compared with the parental cytotoxic moieties (Fig. 5).

Compounds 4a and 5a were used in therapy experiments with nude mice bearing subcutaneous SKRC-52 tumors. The maximum tolerated doses of the 2 prodrugs were determined by dose escalation experiments and found to be 500 nmol/kg for compound 4a (Supplementary Fig. S13) and 50 nmol/kg for compound 5a. The MMAE conjugate 4a, used at a dose of 250 nmol/kg with 10 daily administrations, exhibited a potent antitumor activity effect (P <0.0001 at day 26, compared with the control groups), whereas compound 4b (devoid of the acetazolamide moiety and serving as negative control) did not retard tumor growth (Fig. 6A). In these experimental conditions, no acute toxicity and no significant weight loss was observed for the group of mice treated with the acetazolamide–MMAE conjugate 4a (Fig. 6B). One of 5 mice enjoyed a durable complete response (with no tumor regression for the next 4 months), whereas lesions started to regrow in the remaining 4 mice 1 week after the last injection.

The antitumor efficacy of the PNU-159682 conjugate 5a was investigated at a dose of 25 nmol/kg (Fig. 6C). Also in this case, the conjugate 5a exhibited a potent antitumor effect, whereas the negative control compound 5b did not inhibit tumor growth, confirming the crucial role of the acetazolamide moiety for the potentiation of drug activity. An additional group of animals was injected with equimolar doses of compound 5a and the acetazolamide derivative 3a, leading to a slight decrease in therapeutic activity and tolerability (Fig. 6C). The treatment of mice with PNU-159682 derivatives was associated with higher toxicity compared with the results obtained with MMAE conjugates, but the 5a treatment group completely recovered from a reversible 10% body weight loss (Fig. 6D). At day 40, a second therapeutic cycle of 4 injections was performed, which did not lead to the disappearance of neoplastic lesions. To investigate the reasons for the absence of therapeutic activity in the second treatment cycle, mice were injected with the fluorescently labeled acetazolamide derivative 3a (250 nmol/kg) which exhibited only a faint tumor accumulation (Fig. 7A), in stark contrast to predosing results (Fig. 3B). An immunofluorescence analysis of resected tumors confirmed that cancer cells remained strongly positive for CAIX (Fig. 7B), suggesting that a reduction in vascular permeability after therapy may have impaired drug accumulation in the neoplastic lesions.

Acetazolamide derivatives selectively localize to renal cell carcinoma xenografts in mice, which express CAIX as membrane protein. The interaction of these compounds with CAIX-expressing cells has been investigated at the cellular level, by means of fluorescence microscopy experiments. The latter showed that acetazolamide-bearing conjugates bind to CAIX on the cell membrane without triggering a receptor-mediated endocytosis, in agreement with previous findings (19, 20).

A pool of AAZ-based SMDCs has been prepared by our group in recent years: different types of cytotoxic drugs (DM1, MMAE, or PNU-159682) were attached to acetazolamide via cleavable linkers (i.e., disulfide bond or valine–citrulline peptide). These SMDCs were found to have similar serum half-lives, which were sufficiently high to exhibit therapeutic benefits in vivo. In particular, the new valine–citrulline–bearing compounds 4a and 5a exhibited a half-life of about 7 hours in serum, which is substantially longer than the circulatory half-life of the radiolabeled analogue 1.

While drugs commonly used for the treatment of metastatic kidney cancer (e.g., sorafenib and sunitinib) did not display any detectable activity against SKRC-52 tumors (20), acetazolamide derivatives of MMAE (a tubulin inhibitor) and of PNU-159682 (a nemorubicin metabolite) could suppress tumor growth, sometimes mediating a complete tumor eradication. The removal of the acetazolamide moiety from the linker–payload combinations abrogated therapeutic activity in all experimental systems, confirming that the ligand-based delivery of cytotoxic drugs to the extracellular tumor environment represents a strict requirement for anticancer efficacy. Both compounds 4a and 5a described in this article feature a linker containing a valine–citrulline moiety, which was previously believed to be particularly suited for the intracellular release of drug payloads, due to the action of lysosomal cathepsin B (41). This protein is a primarily intracellular protease that can be secreted extracellularly by dying cells but also by living tumor cells to initiate extracellular proteolytic cascades and to enable tumor cell proliferation (42). Our therapy data suggest that the presence of cathepsin B in the extracellular tumor environment is sufficient for the efficient liberation of cytotoxic drug moieties, which can then diffuse into surrounding tumor cells.

Nude mice do not represent a perfect model for the assessment of anticancer agents for a number of reasons (43), including their lack of T cells. It has recently been shown that certain cytotoxic agents can mediate an immunogenic tumor cell death (44, 45). In addition, certain immunostimulatory agents (e.g., antibody–cytokine fusion proteins, immunologic checkpoint inhibitors) may potently synergize with some cytotoxic agents (46–48).

Despite the fact that a strong suppression of tumor growth was observed both for compounds 4a and 5a, cancer cures were rare. Interestingly, treatment of mice with a second cycle of 5a therapy did not cause a second tumor regression (Fig. 6). Macroscopic and microscopic imaging data suggest that the tumor uptake of acetazolamide derivatives is reduced after therapy, probably as a result of changes in vascular permeability at the neoplastic site, whereas cancer cells remain CAIX-positive. It will be interesting to investigate whether similar findings can be observed with other classes of targeted cytotoxics (e.g., with ADC products).

The potent therapeutic activity observed with compounds 4a and 5a suggests that these products may provide a benefit to patients with kidney cancer, as the majority of clear cell renal cell carcinomas are strongly positive for this target (32–34). The biodistribution profiles of acetazolamide derivatives compares favorably with the ones previously reported for other ligands (23–26). In particular, we were pleased to see that %ID/g values in the tumor were higher than the corresponding values in the kidney (the main organ for drug clearance), even as early as 1 hour after intravenous administration of the product (18). A nuclear medicine trial in patients with metastatic renal cell carcinoma is currently planned for ^99mTc-labeled acetazolamide 1. That study will provide essential information, regarding the tumor:organ ratios that can be expected in patients with cancer and in mouse models of the disease.

D.N. is a board member of, reports receiving a commercial research grant from, has ownership (including patents) in, and is a consultant/advisory board member for Philogen. No potential conflicts of interest were disclosed by the other authors.

Conception and design: S. Cazzamalli, D. Neri

Development of methodology: S. Cazzamalli, A. Dal Corso, D. Neri

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): S. Cazzamalli, A. Dal Corso

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): S. Cazzamalli, A. Dal Corso, D. Neri

Writing, review, and/or revision of the manuscript: S. Cazzamalli, A. Dal Corso, D. Neri

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): S. Cazzamalli, D. Neri

Study supervision: D. Neri

We thank Franziska Bootz, Rémy Gébleux, Dario Venetz, and Jonathan Kiefer for their help with experimental procedures. We also thank Alessandra Villa for CAIX production. The authors acknowledge support of the Scientific Center for Optical and Electron Microscopy ScopeM of the Swiss Federal Institute of Technology ETHZ. We personally acknowledge Justine Kusch for the help with confocal microscopy. Nikolaus Krall is acknowledged for many comments and suggestions.

D. Neri gratefully acknowledges ETH Zürich, the Swiss National Science Foundation (Projects Nr. 310030B_163479/1), ERC Advanced Grant “Zauberkugel,” Kommission für Technologie und Innovation (Grant Nr. 17072.1), Bovena Foundation, and Maiores Foundation for financial support.

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