Anesthesia and Circulating Tumor Cells in Primary Breast Cancer Patients

A Randomized Controlled Trial

Frédérique Hovaguimian, M.D.; Julia Braun, Ph.D.; Birgit Roth Z'graggen, Ph.D.; Martin Schläpfer, M.D.; Claudia Dumrese, Ph.D.; Christina Ewald, Ph.D.; Konstantin J. Dedes, M.D.; Daniel Fink, M.D.; Urs Rölli, M.Sc.; Manfred Seeberger, M.D.; Christoph Tausch, M.D.; Bärbel Papassotiropoulos, M.D.; Milo A. Puhan, Ph.D.; Beatrice Beck-Schimmer, M.D.


Anesthesiology. 2020;133(3):548-558. 

In This Article

Materials and Methods

We used the Consolidated Standards of Reporting Trials recommendations for the reporting of randomized trials.[24] This trial was approved by the local ethical committee (Zurich, Switzerland, registration number PB_2016-01791) and was registered with (NCT02005770,, principal investigator: Beatrice Beck-Schimmer, registration date: December 9, 2013). The study protocol is available on

Trial Design and Participants

This was a parallel-group, randomized, controlled trial conducted at a university hospital (University Hospital of Zurich) and a private clinic (Hirslanden Group, Zurich) in Switzerland. Patients were considered eligible if they were aged 18 to 85 yr, diagnosed with primary preinvasive and invasive breast cancer without distant metastases (stage 0 to III) and scheduled for surgery with or without axillary node dissection. Patients were excluded if they met one of the following criteria: preoperative chemotherapy, possible immune impairment (i.e., autoimmune disease, human immunodeficiency virus, other active cancer, American Society of Anesthesiologists (ASA; Schaumburg, Illinois) Physical Status IV or V), immunosuppressive or chronic opioid therapy, secondary surgery (e.g., for recurrence, reconstruction), or surgery performed under general anesthesia with concomitant regional anesthesia (i.e., epidural catheter, paravertebral blockade, wound infiltration with local anesthetics). Those with a known or suspected hypersensitivity or allergy to anesthetics were considered ineligible. Patients were approached on the day before surgery by research staff, who evaluated eligibility, obtained written informed consent, and enrolled the participants.

Randomization and Blinding

Randomization was performed by research staff using a secure Internet-based system (; accessed April 10, 2018) that stratified patients according to their ASA status and ensured concealment of random allocation. The patients were randomly assigned in a 1:1 ratio to either intravenous anesthesia (propofol group) or inhalational anesthesia (sevoflurane group). Patients remained blinded to their assignment group (standardized induction in both groups), as was the study personnel involved in circulating tumor cell measurements (i.e., outcome assessors did not have access to patient charts).


Anesthesia induction was standardized in both groups using fentanyl (2 to 3 µg/kg), thiopental (4 to 6 mg/kg), and rocuronium (0.6 mg/kg). Patients requiring a rapid sequence induction received 0.9 mg/kg rocuronium instead of 0.6 mg/kg. Further administration of fentanyl during surgery followed a standardized protocol (i.e., 2 µg/kg; total amount, 5 to 10 µg/kg). In the propofol group, anesthesia was maintained using a target-controlled infusion device providing an intravenous propofol dose adjusted to keep Bispectral Index values between 40 and 60; in the sevoflurane group, sevoflurane was provided to keep Bispectral Index values between 40 and 60. Postoperative nausea and vomiting prophylaxis and perioperative analgesia followed standardized protocols that were applied until hospital discharge.


The primary outcome was the number of circulating tumor cells assessed postoperatively by the CellSearch assay (Menarini Silicon Biosystems Inc., USA). Based on immunomagnetic separation, this detection technique uses a magnetic field to isolate ferrofluid-labeled tumor cells of epithelial origin, such as breast cancer cells.[25] This standardized procedure uses antibodies directed against a common molecular signature displayed by circulating tumor cells in breast cancer patients (i.e., the "EpCAM+/CK+/DAPI+/CD45-" signature, where EpCAM indicates epithelial cell adhesion molecule, CK indicates cytokeratin, and DAPI indicates 4',6-diamidino-2-phenylindole). After staining of the isolated cells, circulating tumor cell identification was confirmed by two independent, specifically trained laboratory technicians that were masked to treatment assignment. Identification of circulating tumor cells followed a predefined set of criteria (i.e., morphological features, compatible staining pattern).

Peripheral blood was collected at four different time points, i.e., before the induction of anesthesia (baseline), after surgery but before extubation (0 h), on day 2 (48 h), and on day 3 (72 h) postoperatively. The last measurement was initially planned on day 4 but was rescheduled to day 3 in January 2016 to avoid data loss due to early hospital discharge. This was the only change made to the original trial design.

Secondary outcomes were defined as the maximal circulating tumor cell count value at any time point after surgery (0, 48, and 72 h); circulating tumor cell counts as a binary outcome (using two different cutoff values, i.e., at least 1 and at least 5 circulating tumor cells/7.5 ml blood); and the association between natural killer cell activity and circulating tumor cell counts (see also "Additional Analyses"). Initially, only a cutoff value of a least 5 circulating tumor cells/7.5 ml blood was considered. We added the threshold of a minimum of 1 cell at the time of analysis, because evidence suggested that values as low as 1 circulating tumor cell/7.5 ml blood were associated with poorer prognosis in primary breast cancer patients.[22] No other changes were made to primary/secondary outcomes definitions over the study period.

Statistical Analyses

Sample size calculation was performed using a method accounting for repeated measurements of count data over time.[26] Because evidence on the effect of intravenous or inhalational anesthesia on circulating tumor cell counts was nonexistent, we adopted a conservative approach and assumed that the expected effect size (Cohen's d) between groups would be small (0.3). Thus, assuming a within-subject correlation of circulating tumor cell counts over time of 0.4 and a dropout rate of 10%, we estimated that a total of 232 patients would be required (209 patients without dropout) to detect a difference between groups corresponding to an effect size of 0.3, with a power of 80%, at a significance level of 5% (two-sided). Because the dropout rate was particularly low, the trial ended after enrolling 217 patients.

All analyses were based on intention to treat. Continuous data were expressed as means and standard deviations or as medians and interquartile ranges if distributions were skewed. The primary analysis used a mixed Poisson model with random intercept per patient to account for repeated measurements over time and thus correlated observations within subjects. We opted for this approach because the Poisson model is appropriate for count data (primary outcome of circulating tumor cell counts). The results of the Poisson models are presented as rate ratios, denoting the comparison of circulating tumor cell counts between the two groups. To avoid assuming a linear development of circulating tumor cells over time, time was alternatively included as a factor variable in our model. We also explored the effect of anesthetics on the maximal circulating tumor cell count value at any time point after surgery in additional Poisson models (0, 48, and 72 h).

Because circulating tumor cell detection is usually reported as a binary outcome (i.e., positive vs. negative endpoint using a cutoff value of at least 1 or at least 5 circulating tumor cells/7.5 ml blood), circulating tumor cell count data were dichotomized and further assessed using a mixed logistic regression model with random intercept per patient. Finally, models were adjusted to account for tumor-related and perioperative factors presumed to affect circulating tumor cell counts (i.e., tumor size, tumor type, and overall opioids consumption, all preplanned).

All statistical analyses were conducted in R, version 3.6.1. Two-sided tests were performed, and a level of significance of 0.05 was used.

Additional Analyses

Because of the interplay between natural killer cell cytotoxic activity and tumor growth, we also assessed natural killer cell activity (i.e., apoptosis rate induced in tumor cells) in a preplanned, exploratory, in vitro study nested within this trial. Natural killer cell-induced apoptosis was evaluated in a subgroup of patients randomly selected from the study data set. For each patient, natural killer cell activity was assessed at a single, predefined time point, i.e., when circulating tumor cell counts reached their maximal value. The association between natural killer cell-induced apoptosis rate and circulating tumor cell count was then assessed using linear regression analysis.

Natural killer cell-induced apoptosis rate and necrosis rate were determined in vitro by measuring target cell killing of the K562 tumor cell line (human chronic myelogenous leukemia, ATCC, CCL-243).[27,28] Patients blood samples were collected in EDTA-coated vials. Buffy coats (Blutspende Zurich, Switzerland) were used as controls. Peripheral blood mononuclear cells of both patient samples and buffy coats were isolated by Ficoll-Hypaque density gradient centrifugation and stored in liquid nitrogen. For determination of natural killer cell activity, peripheral blood mononuclear cells were thawed and coincubated with K562 for 24 h at 37°C with 5% CO2 in 10% human serum/RPMI medium. An effector (natural killer cells)-to-target cell (K562 cells) ratio of 1:1 was used. All cells were then washed in phosphate-buffered saline and stained in 2% bovine serum albumin in phosphate-buffered saline for 25 min at 4°C using the following panel: CD3-APC (lymphocyte staining; Biolegend, United Kingdom), dilution of 1:100; CD 56-PE (natural killer cells staining; Biolegend), dilution 1:100; and CD16-FITC (FcγRIIIA staining, which is essential for cellular cytotoxicity, expressed on the surface of a subset of monocytes; Biolegend), dilution 1:200. After a washing step in annexin V binding buffer, the cells were simultaneously stained with annexin-PerCPCy5.5 for staining of apoptotic cells (Biolegend) at a dilution of 1:20 and Zombie-NIR for staining of necrotic cells (Biolegend) at a dilution of 1:500.

Zombie-NIR-stained K562 boiled for 5 min at 80°C or annexin V-stained apoptotic K562 and treated for 24 h with 10 mM benzamide were used as positive controls for cytotoxicity. Unstained K562, unstained patient peripheral blood mononuclear cells, and unstained peripheral blood mononuclear cells from buffy coats served as negative controls. Cell analysis was performed using the spectral analyzer SP6800 (Sony Biotechnology, United Kingdom).[29]