Epidermal growth factor receptor inhibitors (EGFRIs)have demonstrated improved overall survival in patients with non-small cell lung cancer, pancreatic cancer, and colorectal cancer; however, their use is associated with dermatologic reactions of varying severity. The similar spectrum of events observed with monoclonal antibodies and tyrosine kinase inhibitors suggests such toxicities are a class effect. While such reactions do not necessarily require any alteration in EGFRI treatment, being best addressed through symptomatic treatment, there is limited evidence on which to base such therapies. In October 2006, at an international and interdisciplinary EGFRI dermatologic toxicity forum, the underlying mechanisms of these toxicities were discussed and commonly used therapeutic interventions were evaluated. Our aim was to reach a current consensus on management strategies. A three-tiered, EGFRIfocused toxicity grading system is suggested for the purposes of therapeutic decision making, and as a framework on which to build a stepwise approach to intervention. This approach to successful management is specifically tailored to accurately categorize dermatologic toxicity associated with EGFRIs, and can be easily applied by all health care professionals. The goal is to maximize quality of life in patients who are being treated with these agents-many of whom will be on these drugs for several months or even years. The Oncologist 2007;12:610 -621 Disclosure of potential conflicts of interest is found at the end of this article.

Background: Circulating tumor cells (CTC) associated with solid tumors are being studied for their diagnostic and prognostic value. In patients with metastatic tumors, CTC presence in the blood has been putatively associated with short survival. Since blood collection is relatively non-invasive, CTC molecular analysis opens up the possibility of monitoring genotypic changes during cancer treatments. Unfortunately, CTCs are not present in large numbers, often at rates as low as one cell per 106-107 leukocytes. Thus, to perform genotypic biomarker analysis on CTCs, methodologies must be developed to using highly specific and sensitive technologies and an enrichment step to increase analytes to detectable levels. Methods: We developed a methodology for detecting mutations in multiple oncogenes and chemotherapy resistance genes in non-small cell lung cancer (NSCLC) CTC specimens using high-throughput matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF MS) single nucleotide polymorphism (SNP) analysis (MASSarray; Sequenom, Inc.) to determine cancer-associated genetic mutations in lung cancer specimens. This system allows for up to 10 different somatic mutations to be assayed per well in a 384-well format; requires very little DNA; can be used with whole genome amplified (WGA) DNA; and is sensitive enough to use with small samples such as core needle biopsies (CNB), fine needle aspirates (FNA), and CTCs. We developed a lung cancer assay panel of 13 genes/135 mutations (including AKT1, BRAF, CTNNB1, EGFR, ERBB2, KRAS, MEK1, NRAS, PIK3CA, PIK3R1,PTEN, and STK11) to test for somatic mutations in genes representing multiple pathways known to be involved in lung cancer. All assays can detect a mutation in < 25% of a sample. Results: In the effort to analyze CTCs, we first analyzed 57 NSCLC cell lines with known mutations and confirmed known mutation status. Next, we successfully analyzed DNA from 90 frozen and matched FFPE NSCLC resected tissues. Analysis of unamplified and matched WGA cell line DNA quantity CNB and FNA equivalents gave the same mutational status results. Moving to CTC equivalents, we successfully analyzed cell line DNA and matched WGA DNA equivalents of 100–1000 cells with known EGFR L585R and KRAS G34A mutations and negative control DNA (negative for all assays). Next, WGA methodology for direct CTC cell lysate DNA amplification was developed using CTC cell number equivalents (3 – 200 cells) obtained from a typical clinical blood sample CTC preparation. Then, we directly compared unamplified and matched amplified CTC cell equivalents (50, 100, and 200 cells). Analysis of both unamplified and amplified CTC cell equivalents reported identical mutation status results. We have applied this methodology to spiked blood sample and clinical blood sample CTC fractions. Thus, we demonstrated that we are able to study mutations in multiple genes using small amount of DNA from CTC cell numbers in a high-throughput manner. Conclusion: We developed a robust method for accurately determine cancer-associated genetic mutations in NSCLC CTC cell number equivalent lysates using MALDI-TOF MS SNP analysis which can be applied to better understand the molecular characteristics of lung cancer during treatment and progression. As additional clinical NSCLC CTC samples are collected, we will continue applying this methodology to assess CTC mutation status as potential diagnostic and/or prognostic markers.