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Felton et al. J Cancer Metastasis Treat 2018;4:51 I http://dx.doi.org/10.20517/2394-4722.2018.39 Page 3 of 9

tion of tumor tissue directly into the wall of the colon have the potential to be more representative of human
[14]
metastasis .

We sought to determine if mice could serve as a model to explore regional differences in the location of cancers
within the colon. To seek such differences in the growth and progression of colon neoplasia in mouse models,
we first determined the location of colon tumors in mice treated with a colon-selective carcinogen or with a
genetic predisposition to intestinal neoplasia. Next, based on our initial findings, we compared the procliv-
ity of human colon cancer cells to grow and invade the proximal vs. distal colon of immune-deficient mice.

METHODS
Analysis of the distribution of colon neoplasia in our published studies of chemically-induced
carcinogenesis in mice
To assess the regional distribution of murine colon cancer, we re-analyzed data from our published and
unpublished murine colon cancer studies conducted from 2006 through 2018 [15-19] . During this interval, we
had treated 10- to 23-week-old male mice on a variety of genetic backgrounds with weekly intraperitoneal
injections of 7.5 mg azoxymethane (AOM)/kg body weight for 4 weeks. In C57BL/6 mice that are resistant to
[20]
AOM treatment alone , we supplemented the drinking water with 2.5% dextran sodium sulfate (DSS) for
5 days. We euthanized mice 20 weeks after the first AOM injection. An investigator masked to mouse geno-
type and treatments measured tumor number and size, and tumors were characterized as adenomas or ad-
enocarcinomas based on size, contour, and color. A senior pathologist classified colon tumors as adenomas
[21]
or adenocarcinoma based on consensus recommendations .

Surgical induction of colon neoplasia
Cell culture
We purchased authenticated HT-29 cells from American type culture collection (ATCC). HT-29 cells were
grown in McCoy’s 5A medium (Life Technologies) supplemented with 10% FBS. We grew cells in a humidi-
fied incubator at 37 ºC with 5% CO and passaged weekly at subconfluence after trypsinization. We suspend-
2
6
ed cells in DPBS (50 × 10 cell/mL) containing 10 µmol/L Y27632 and 50% Matrigel.
Animals
All animal studies were conducted at the Baltimore VA Hospital Animal Facility and our laboratory in the
Bressler Research Building at the University of Maryland School of Medicine. All surgical procedures were
approved by the University of Maryland School of Medicine Institutional Animal Care and Use Committee
under the Office of Animal Welfare Assurance. The Research and Development Committee at the Baltimore
nu
VA also approved animal studies. We used 11- to 14-week old male Nu(NCr)-Foxn1 (nude) mice and NOD.
scid
Cg-Prkdc Il2rg Tim1Wji /SzJ (NSG) mice, obtained from both the University of Maryland Veterinary Resources
and Jackson Laboratories (Bar Harbor, ME).

Surgical technique - laparotomy
In a biosafety cabinet (BSL2) we anesthetized mice with continuous vaporized isoflurane for general anes-
thesia and performed laparotomy and cell injections with the mice on a warming pad. After confirming a
sufficient level of anesthesia by a toe pinch, we positioned mice prone. For corneal protection, we applied
lubricant (Major Pharmaceuticals LubriFresh P.M Ophthalmic Ointment, Livonia, MI) to each eye. We dis-
infected the mouse’s upper back with an alcohol swab and administered buprenorphine SR (concentration
0.3 mg/mL, dose 0.05-0.1 mg/kg body weight diluted 1:9 with sterile 0.9% saline) or carprofen (concentration
50 mg/mL, dose 5 mg/kg body weight diluted 1:9 with sterile 0.9% saline) subcutaneously for analgesia. We
then placed mice supine and, if necessary, clipped the anterior abdominal hair. After skin preparation with
alcohol, to provide local anesthesia we injected mice subcutaneously with 0.25% bupivacaine (concentration
2.5 mg/mL, dose 0.1 mL diluted 1:2 with sterile 0.9% saline) along the planned midline laparotomy site. Next,