BMC Medicine

BioMed Central

Research article

Open Access

Collagen reorganization at the tumor-stromal interface facilitates
local invasion
Paolo P Provenzano*1,2, Kevin W Eliceiri2, Jay M Campbell2,
David R Inman1, John G White2 and Patricia J Keely*1,2
Address: 1Department of Pharmacology, University of Wisconsin, Madison, WI, USA. and 2Molecular Biology Program, Laboratory for Optical andComputational Instrumentation, University of Wisconsin, Madison, WI, USA.
Email: Paolo P Provenzano* - paolo.provenzano@gmail.com; Kevin W Eliceiri - eliceiri@wisc.edu; Jay M Campbell - jmcampbe@wisc.edu;
David R Inman - drinman@wisc.edu; John G White - jwhite1@wisc.edu; Patricia J Keely* - pjkeely@wisc.edu* Corresponding authors

Published: 26 December 2006
BMC Medicine 2006, 4:38

doi:10.1186/1741-7015-4-38

Received: 18 June 2006
Accepted: 26 December 2006

This article is available from: http://www.biomedcentral.com/1741-7015/4/38© 2006 Provenzano et al; licensee BioMed Central Ltd.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0),
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract
Background: Stromal-epithelial interactions are of particular significance in breast tissue as misregulation ofthese interactions can promote tumorigenesis and invasion. Moreover, collagen-dense breast tissue increases therisk of breast carcinoma, although the relationship between collagen density and tumorigenesis is not wellunderstood. As little is known about epithelial-stromal interactions in vivo, it is necessary to visualize the stroma
surrounding normal epithelium and mammary tumors in intact tissues to better understand how matrixorganization, density, and composition affect tumor formation and progression.
Methods: Epithelial-stromal interactions in normal mammary glands, mammary tumors, and tumor explants inthree-dimensional culture were studied with histology, electron microscopy, and nonlinear optical imaging
methodologies. Imaging of the tumor-stromal interface in live tumor tissue ex vivo was performed withmultiphoton laser-scanning microscopy (MPLSM) to generate multiphoton excitation (MPE) of endogenous
fluorophores and second harmonic generation (SHG) to image stromal collagen.
Results: We used both laser-scanning multiphoton and second harmonic generation microscopy to determine
the organization of specific collagen structures around ducts and tumors in intact, unfixed and unsectioned
mammary glands. Local alterations in collagen density were clearly seen, allowing us to obtain three-dimensional
information regarding the organization of the mammary stroma, such as radiating collagen fibers that could nothave been obtained using classical histological techniques. Moreover, we observed and defined three tumorassociated collagen signatures (TACS) that provide novel markers to locate and characterize tumors. In particular,
local cell invasion was found predominantly to be oriented along certain aligned collagen fibers, suggesting thatradial alignment of collagen fibers relative to tumors facilitates invasion. Consistent with this observation, primarytumor explants cultured in a randomly organized collagen matrix realigned the collagen fibers, allowing individual
tumor cells to migrate out along radially aligned fibers.
Conclusion: The presentation of these tumor-associated collagen signatures allowed us to identify pre-palpable
tumors and see cells at the tumor-stromal boundary invading into the stroma along radially aligned collagen fibers.
As such, TACS should provide indications that a tumor is, or could become, invasive, and may serve as part of astrategy to help identify and characterize breast tumors in animal and human tissues.

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Background
Tissue microenvironments play an important role inmaintaining normal cell behavior [1-3]. Moreover, type I
collagen is a prevalent component of the stromal extracellular matrix; its expression being spatially and temporally
regulated during mammary ductal formation, suggesting
it plays important roles in development [4]. Consistent
with this idea, decreasing the levels of α2β1 integrin, a primary type I collagen receptor, disrupts mammary epithelial tubulogenesis in vitro [2] and alters branchingmorphogenesis in vivo [5], respectively. Furthermore,inappropriate stromal-epithelial interactions can promotetumorigenesis [6-8], and in breast cancer, metastatic epithelial cells migrate in direct contact along stromal collagen fibers [9]. The importance of studying stromalinteractions in breast tissue is further reinforced by thefact that patients with collagen-dense breast tissue possessa greater than fourfold increased risk of breast carcinoma[10,11]. Although the mechanisms mediating the effectsof the extracellular matrix (ECM) on breast carcinoma
development in vivo are largely unknown, contributing
factors may be adhesion mediated signaling and mechanical signals imparted on mammary epithelial cells fromsurrounding type-I collagen-rich stroma, either directly or
across basement membrane proteins. One important stepto elucidating these signaling interactions is to determine
the organization of the collagenous stroma surrounding
both normal mammary glands and tumors within intact
tissue so as to better understand the cell-matrix interaction and how matrix organization, density, and composition affect tumor formation and progression.
Nonlinear microscopy techniques such as multiphoton
laser-scanning microscopy (MPLSM) and second harmonic generation (SHG) provide powerful tools to image
cellular autofluorescence and extracellular matrix structure in intact tissues [12-15]. Both techniques are well
suited for high-resolution in vivo imaging, and second
harmonic generation is particularly adept at imaging collagen structure. Specifically, multiphoton microscopy
results from the nonlinear excitation of molecular fluorescence and can produce images deep into thick tissues[16,17], while SHG signals depend on non-linear interactions of illumination with a non-centrosymmetric environment (e.g. fibrillar collagen) that can provide
submicron resolution [13-15]. The most commonly utilized multiphoton process is two-photon excitation (2PE)of fluorescence, in which two low-energy (usually nearinfrared) photons simultaneously excite a fluorophore,
which later decays to produce a single fluorescent photonof lower energy than the corresponding one-photon (halfwavelength of 2PE) excitation [13,18,19]. In this 2PE
process the fluorescence is dependent upon the square of
the intensity (see Methods), producing optical sectioningthat makes it equivalent to confocal imaging in terms of

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restricting excitation to the plane of focus, but facilitates amuch greater effective imaging depth and better cell viability [16,20]. SHG imaging, on the other hand, does notarise from an absorptive process, but instead the laser field
suffers a nonlinear, second-order, polarization when passing through certain ordered structures resulting in a coherent signal at exactly half the wavelength of the excitation[21]. Great utility arises from the fact that MPLSM and
SHG can be implemented simultaneously in live tissue toprovide complementary information and a powerfulexperimental and diagnostic tool.
The purpose of this study was to characterize collagenmorphology in intact tissues so as to understand the structure-function relationship of epithelial- and tumor-stromal interactions in the mammary gland with particularemphasis on local tumor cell invasion during carcinoma
progression. We used both MPLSM and SHG imaging, inconjunction with additional correlative microscopy techniques, to detect differences in local collagen density nearnormal glands and mammary tumors, and identify distinct collagen fiber organization around tumors, withcharacteristic collagen structures such as radially aligned
collagen fibers associated with tumor-cell invasion. Identification and characterization of these collagen signatures
sheds insight into the process of tumor cell invasion, andmay serve a diagnostic capacity for determining the invasive potential of tumors.

Methods
Mouse mammary tissues and tumors
This study was approved by the institutional animal use
and care committee and meets N.I.H. guidelines for animal welfare. To study non-tumor bearing mammary
glands, tissue was obtained from B6129SF2/J mice orCol1a1tmJae mice (The Jackson Laboratory, Bar Harbor,
ME, USA). To study tumor-stromal interactions in intact
tissue, two mouse breast tumor models were utilized:
MMTV-Wnt-1 (colony founder mice provided by Dr.
Caroline Alexander, University of Wisconsin, Madison,
WI, USA) and MMTV-polyoma middle-T (abbreviated
PyVT following the Jackson Laboratory title but is also
commonly abbreviated as PyMT or PyV-MT; colony
founder mice originally obtained from Jackson Laboratory were provided by Dr. Amy Moser, University of Wisconsin, Madison, WI, USA).
Tumor explants and collagen gel culture
To study tumor-mediated collagen reorganization andtumor cell invasion in vitro, tumor explants wereobtained and cultured in a manner similar to a previous
report by Friedl and co-workers [22]. Small pieces oftumor were harvested from the central region of palpable
PyVT tumors (that were confirmed by histology) with a 3
mm biopsy punch and were cultured in type I collagen

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gels. Following removal, tumors were rinsed in DMEM
containing penicillin/streptomycin/fungizone solution(Cellgro, Herndon, VA). A single tumor explant was thencultured within a 2.0 mg/ml collagen gel (8.0 mg/ml rattail collagen solution (BD Biosciences, San Jose, CA) neutralized with 100 mM HEPES in 2 × PBS). Following Gelpolymerization for 1 hr, the tumor explant containing collagen gels were released from the culture dish and floatedin DMEM containing penicillin/streptomycin solution
supplemented with 10% heat inactivated FBS. Imaging
was performed on live (non-fixed) cells in intact threedimensional collagen gels.
Multiphoton microscopy and second harmonic generation
Multiphoton excitation (MPE) with MPLSM allows imaging of endogenous fluorophores from deep inside live
biological tissues with the fluorescence excitation primarily restricted to the plane of focus due to a quadratic
dependence on the laser light intensity and a low probability of multiple low-energy photons being absorbed outside the focal plane [16,17,23]. For the case of a pulsed
laser excitation, the time averaged fluorescent intensity isa function of the molecular cross-section δ2(λ) and thesquare of the laser intensity, I(t)2, and can be expressed as[23,24]:P2 ⎡ (NA)2 ⎤If ,p (t) = δ 2 ave ⎢ π⎥
τ p fp ⎢⎣ hcλ ⎥⎦2(1)

In Equation 1, δ2 is defined as a molecular cross sectionthat represents the dependence for the probability of twophoton excitation on the square of photon density, P is
the laser power, τp is the laser pulse width, fp is the laser
repetition rate, NA is the numerical aperture, h is Planck's
constant, c is the speed of light, and λ is the wavelength.
In contrast to multiphoton excitation, which obeys thefundamental physical relationship of energy loss following excitation, SHG is a conserved polarization processthat follows the relationship [21,25,26]:
P = χ(1) * E + χ(2) * E * E + χ(3) * E * E * E + …

(2)

where the polarization (P) and electric field (E) are vectors, and the nonlinear susceptibilities, χ(i), are tensors.
Therefore, SHG arises from the laser field suffering a conserved nonlinear, second-order, polarization when passing through non-centrosymmetric ordered structures thatis described by term 2 of Equation 2.
For MPE and SHG imaging of live unfixed, intact (not sectioned), non-stained glands, and tumor explants withincollagen gels, as well as hematoxylin and eosin stained
histology slides, we used an optical workstation [27] thatwas constructed around a Nikon Eclipse TE300. For live
tissue imaging, twenty mammary tissues including ninefrom the Col1a1tmJae strain (three each of wild type, heterozygous and homozygous), and tumors from Wnt-1 (n =10, plus wild type controls) and PyVT (n = 20) mice were
harvested and live tissue maintained in buffered media at
37°C. All tissues were imaged immediately following tissue harvest and a Ti:sapphire laser (Spectra-Physics-Millennium/Tsunami, Mountain View CA) excitation source
producing around 100 fs pulse widths and tuned to 890–
900 nm was utilized to generate both multiphoton excitation (cellular autofluorescence from FAD) and SHG. The
beam was focused onto the sample with either a Nikon
40× Plan Fluor oil-immersion lens (N.A. = 1.4) or a Nikon
60× Plan Apo water-immersion lens (N.A. = 1.2). All SHG
imaging was detected from the back-scattered SHG signal[26], and the presence of collagen confirmed in our tissues using fluorescence lifetime imaging microscopy, or
FLIM [12], on the same optical workstation, as the SHG
from collagen has no lifetime. Additionally, due to thefundamental differences between MPE and SHG signals,
filtering can separate the emission signals. Using a 464
nm (cut-on) long pass filter, MPE was discriminated fromthe total emission while a 445 nm narrow band pass filters was used to separate SHG (filters from TFI Technologies, Greenfield, MA). For 3D imaging in intact tissues, 2D(x-y) images were acquired at various serial depths (z)into the samples.
Acquisition was performed with WiscScan [28] a software
acquisition package developed at LOCI (Laboratory for
Optical and Computational Instrumentation, Universityof Wisconsin, Madison, WI, USA). Image analysis for
combined MPE-SHG was performed with ImageJ [29] andVisBio [30] software. Using ImageJ, differences in collagendensity were quantified by measuring the area of collagensignal following density slicing from a constant threshold,and local and mean intensity were measured within these
normalized areas. For TACS-1 image analysis additionalsurface rendering plug-ins for ImageJ were utilized. For
TACS-2 and -3, ImageJ was used to quantify the collagenfiber angle relative to the tumor. The tumor boundary wasdefined and the angle relative to the tangent of tumor
boundary was measured every 10 microns.
Histology and electron microscopy
For histology, formalin-fixed paraffin-embedded samples
from eight B6129SF2/J mice and eight Col1a1tmJae mice
were sectioned and stained for hematoxylin and eosin, trichrome, and picrosirius red using standard techniques.
Additionally, all tissues imaged with MPLSM were subsequently fixed and processed for histology to confirm thepresence of tumors and characterize the tumor morphology. Sample preparation for scanning and transmission
electron microscopy (SEM and TEM, respectively) was

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performed by fixing whole mammary glands in 2.5% formaldehyde/2.5% glutaraldehyde in 0.1 M sodium
cacodylate buffer for 1 hr at room temperature (RT), after
which sample were placed in fresh fixative overnight at
4°C. Samples were then washed in cacodylate buffer andpostfixed in 1.5% osmium tetroxide at RT for 1.5 hrs.
Samples (SEM n = 8 glands and TEM n = 6 glands) from
B6129 mice were again washed in buffered solution anddehydrated, fractured, critical point dried, sputter coated,
and imaged with SEM, or stained, dehydrated and cleared,
embedded, and then sectioned for imaging with TEM[31].

Results
The normal mammary gland
Examination of the ECM surrounding mammary epithelial cells revealed that the majority of the stroma is fibrillar
collagen, which was seen surrounding epithelial cells (Figure 1). Figure 1A, in which picrosirius red (a selective
staining agent for collagen) was used indicates the presence of collagen around the mammary ductal structure(Figure 1A,B). Observations at high magnification with
SEM showed collagen fibers wrapping around the ductal
structure in an organized manner surrounding the epithelial cells (Figure 1C), in a fashion similar to that seen in
Figure 1A. Additionally, collagen can be seen wrapping
individual cells in multiple directions (Figure 1C–E), suggesting a role for collagen as a containing and anchoring
structure. Of further importance, higher magnification
images with both SEM (Figure 1F,G) and TEM (Figure 1H)
confirmed that the collagen fibers are composed of fibrillar collagen (Figure 1F; as indicated by their rod-like structure and presence of the ~67 nm banding pattern [32])that are adjacent to the epithelial cells (Figure 1G,H).
These data, in combination with published works indicating an important interaction between the stroma and theepithelial cell (see [4,5,33]), could suggest that these fibers apply physical restraint and mechanical signals acrossa thin basement membrane (<200 nm), or that the thinmesh-like basement membrane may contain some small
gaps or membrane spanning cell processes and therefore
may not completely isolate the epithelial cell; resulting in
micro-regions of direct collagen-epithelial cell interactions [34]; or, most likely, a combination of both scenarios.

Although histology and electron microscopy provide valuable and detailed information regarding the composition and morphology of the epithelial-stromalinteraction, these techniques are destructive to the sample
and the capability for three-dimensional imaging is limited. Therefore, we have sought non-destructive imaging
techniques (e.g. MPE/SHG) that allow imaging in four
dimensions (x, y, z, and time). Combined MPE-SHG produced clear images of collagen (Additional file 1) as well

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as endogenous fluorophores, such as NAD(H) and FAD(Additional file 1, part C), in intact non-treated tissues[13,15,21]. Application of MPE/SHG to intact, non-fixed,
non-stained mammary gland captured the morphology ofthe fibrous stroma seen in Figure 1. For example, images
captured "above" the gland (Figure 2A: a and 2A: g) clearly
showed the presence of wavy (crimped) collagen as wellas the presence of taut (straight) fibril bundles (Figure 2A:
a,e,f,g). Taut fibers showed a periodicity of ~250 nm (Figure 2A: c), consistent with previous reports for SHG resolution [14], which may represent an approximately
fourfold super-periodicity of the basic ~67 nm banding
pattern of collagen fibrils noted with SEM (Figure 2A: d).
Additionally, collagen was observed wrapping around theepithelial duct (consistent with Figure 1A,C), as well asradiating away from the duct (Figure 2A: b). Furthermore,in addition to detecting unique features from normal
mammary glands, examination of the epithelial-stromalinteraction in Wnt-1 mice [35], which possessed ductal
hyperplasia (Figure 2B: a–c), revealed abnormal mammary duct development with irregular collagen organization (Figure 2B: b,c). The ability to clearly discern thesepathologic changes in tissues that were not fixed, sectioned, or stained (Figure 2B: d–f) validates the application of MPE/SHG imaging to detect normal mammary
gland and collagen structure as well as abnormalities inmammary ducts and collagen distribution in situ (Figure
2B: f).
Detecting dense mammary tissue
High breast tissue density (due to increased collagen;[36]) is one of the single largest risk factors for developing
breast cancer [37], yet the molecular mechanisms behind
this high risk are not known. One reason for this deficit
has been the lack of adequate animal model systems forstudying the effects of increased collagen density in vivo.
As such, we sought to identify an animal model system
possessing collagen-dense breast tissue. Analysis of the
col1a1tmJae mouse model [38] revealed such a system.
These mice possess a type I collagen mutation in the α1(I)chain, making them resistant to human collagenase,resulting in fibrosis of the skin and uterus due to excessive
collagen accumulation [38]. However, increases in collagen deposition in the mammary glands of these mice havenot been previously described. Analysis with standard histology clearly revealed increased collagen surrounding the
mammary ducts in both homozygous and heterozygous
female mice regardless of parous status (Figure 3A vs 3C;
Additional file 2). By utilizing histology and imaging
hematoxylin and eosin (H&E) stained sections with
MPLSM (SHG signal did not exist after formalin fixation;
data not shown), the presence of increased collagen was
confirmed (Figure 3B vs 3D), as well as an apparent hyperplasia in Col1a1 mice (Figure 3C,D; Additional file 2).
Interestingly, this hyperplasia was associated with inva-

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Figure 1morphology and the epithelial-stromal interaction in the mammary gland
Collagen
Collagen morphology and the epithelial-stromal interaction in the mammary gland. (A) Picrosirius red staining(red), a selective collagen stain, indicating the primary stromal component is collagen (bar = 50 μm). (B) Low magnification (bar= 10 μm) TEM image of a mouse mammary duct (transverse section) showing the organization of epithelial (e) and myoepithelial (me) cells outside the lumen (L) in close association with the collagenous stroma (s), with myoepithelial cells not completely
separating the stroma from the epithelial cells. (C) SEM image of the ductal end. (D) SEM image of collagen fibers interactingwith ductal epithelial cells. (E) SEM image of collagen bundles wrapping around the cell in multiple directions. (F) Collagen fibrils
possessing a rod-like structure with the characteristic ~67 nm banding pattern validating the presence of fibrillar collagen inproximity to epithelial cells. (G) SEM image of collagen fibrils immediately adjacent to the cell surface. (H) TEM image of collagen fibrils (col) next to the epithelial cell (e).

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Figure 2
MPLSM/SHG
imaging reveals native collagen structure in living mammary gland
MPLSM/SHG imaging reveals native collagen structure in living mammary gland. A: (a) MPE/SHG image at the"top" of the mammary duct, showing both wavy and taut (straight) collagen structures as well as endogenous fluorescencefrom FAD in stromal cells (most likely fibroblasts and immune cells). (b) MPE/SHG image of a mammary duct demonstrating
collagen wrapped around the duct as well as radiating out from the duct (as can be seen predominantly at the top of the micrograph near the end of the duct). (c) Enlarged section from (a) showing some aspects of collagen fibril structure that resemble
the standard banding pattern seen in collagen fibrils from connective tissue. (d) SEM of mouse tendon mouse tendon demonstrating the ~67 nm banding pattern characteristic of collagen fibrils. (e) Correlative SEM image of collagen surrounding ductal
epithelial cells showing both wavy (upper arrow) and taut (lower arrow) fibers that match the collagen structures obtainedwith MPE/SHG imaging. (f) MPE/SHG image of the region near the nipple in tissue demonstrating straightened collagen fibers
radiating from the central ductal structure. (g) MPE/SHG image "above" the mammary duct showing both wavy and taut fiber
structures. Note: all mammary tissues in A are from the B6/129 strain, which serves as the background strain for col1a1 mice.
B: (a) Whole mount analysis from wild type control (top) and Wnt-1 mice (bottom), which display hyperplasia compared towild type glands. (b,c) Histology of mammary glands from wild type control animals (b; H&E) and hyperplastic abnormal ducts
from Wnt-1 mice (c; Masson's Trichrome). (d,e) MPLSM/SHG imaging of wild type control mammary duct demonstrating aductal end (d) and branching (e). (f) MPLSM/SHG imaging of a mammary duct from a Wnt-1 mouse showing hyperplasia of theductal structure and increased deposition of disorganized collagen. Note: All MPE/SHG images are from live intact tissues thatare not fixed, sectioned, or stained; scale bar for MPE/SHG images equals 25 μm; scale bar for histology images equals 50 μm.

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sive-looking epithelial cells that resemble an epithelialmesenchymal transition at the ductal end (Figure 3C,D).
Therefore, in an approach opposite to that taken by Jain
and co-workers [39] where a decrease in collagen in vivo
was detected with SHG, we detected increased collagen inlive intact mammary glands. Analysis of glands fromcol1a1 heterozygous and homozygous mice, and associated wild type littermates, noticeably revealed increased
collagen surrounding epithelial ducts from transgenic
mice (Figure 3E–H), demonstrating our ability to detect
differences in collagen density. Furthermore, collagen was
locally dense adjacent to (wrapped around) the epithelium, as well as increased in the space extending from theduct resulting in a 2.53- (± 0.24) and 2.79- (± 0.34) (mean± SEM) fold increase in the area of collagen signal, as wellas increased signal intensity around the duct, for heterozygous and homozygous col1a1tmJae mice relative to wild
type animals, respectively. Therefore, the col1a1 mousemodel appears to be a promising candidate for studying
the effects of increased collagen density in normal as wellas transformed epithelial cells as appropriately crossed
heterozygous col1a1 mice are capable of forming tumors
without inhibition from the collagenase resistant matrix(P.P.P. and P.J.K unpublished observations). Moreover,
we could discern changes in density and obtain images
from deep (maximum Depth: 440 nm) within intact
mammary tissue (Figure 3H; Additional file 4) without
fixing, sectioning, or staining the tissues, suggesting that
this approach could have wide application in live animalor tissue models for understanding stromal changes associated with pathologic conditions.
Tumor-associated collagen signatures (TACS)
Tumor-associated collagen signature-1
Analysis of tumor bearing Wnt-1 mice, which progress
through hyperplasia, mammary adenocarcinoma, andinvasive (metastatic) ductal carcinoma, revealed multiple
epithelial clusters, containing hemorrhagic regions, intermixed and surrounded by increased collagenous stroma(i.e. desmoplasia), consistent with previous reports [35].
Importantly, the presence of increased collagen allowed
us to identify pre-palpable tumors (Figure 4A: a–c, Additional file 5; and confirmed with histology – data notshown) by the existence of what we are classifying as oneof three tumor-associated collagen signatures (TACS).
Namely, TACS-1: the presence of dense collagen (Figure
4A: a), indicated by increased signal intensity (see surface
map in Figure 4A) at a region near the tumor (Figure 4A:a; Additional file 5) that served as a reliable hallmark forlocating small tumor regions. Although increased collagen has been reported near tumors, the structure of suchcollagen has previously been largely unknown, and the
additional focal localization shown in Figure 4A indicatesincreased local areas of high density within the globally
increased collagen concentration surrounding tumors.

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However, it is unclear whether: (1) a pre-tumor dense
region is present that serves to stimulate tumor formation,(2) the dense region of collagen is pulled into a grouped
cluster through increased contraction of an epithelial
tumor mass or motile cells at the tumor boundary, or (3)fibroblast activation results in increased local collagendeposition. Further analysis into this fundamental characteristic should provide useful information on tumor initiation and progression.
Tumor-associated collagen signatures-2 and -3
As the size of the tumor increased, we identified a second
collagen signature, TACS-2: "taut" collagen fibers
stretched around the tumor (Figure 4B). This collagenmorphology likely arose from stretching of the stroma
due to tumor growth, which may act to constrain portionsof the tumor (i.e. compressive restraint) as well as provide
a stretch induced tensile stress in expanded fibrils (andlarger resistance to cell contraction in the stroma) that
stimulates and activates fibroblasts. Evidence for tumor
restraint in Wnt-t mice can be seen in Figure 4B, a–c,
where collagen fibers are stretched around a relatively
smooth tumor boundary as indicated by the fact that thefiber angle is primarily distributed tangentially (0° relative to the tumor boundary) along the tumor boundary(see histogram in Figure 4B).

In regions of tumor masses that are undergoing growthand invasion (Figure 4C: a–c), a third tumor-associatedcollagen signature, TACS-3, was identified: collagen fibersaligned normal to tumor boundary regions that display anirregular shape – indicative of local invasion through collective epithelial cell migration [22,40]. This invasive
tumor morphology was seen in regions of tumors wherecollagen fibers are primarily aligned in the direction ofcell invasion (see histogram in Figure 4C, in which theangle of the collagen fibers relative to the tumor boundary
distributes around 90°). Furthermore, at regions where
TACS-3 is noted, we observed groups of cells advancing
from the tumor boundary in a collective manner thatappear to be undergoing collective invasion [22,40,41], as
well as individual invading cells that may relate to single
cell migration similar to what has been observed alongcollagen fibers in an in vivo xenograft model [9].
To further investigate the behavior of the third collagensignature in a more invasive and metastatic cancer model,
we utilized the well established and characterized PyVT
mouse model. This model bears resemblance to many
aspects of human cancer, is reliably invasive and metastatic following defined, progressive, and reliable histological grades from hyperplasia to adenoma, and then to
early and late carcinoma, and therefore provides a goodmodel for studying human disease [42]. Tumor cell
behavior and collagen morphology in this model were

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Figure 3
MPLSM/SHG
imaging detects increased collagen density in the mammary gland
MPLSM/SHG imaging detects increased collagen density in the mammary gland. (A) H&E stain of wild type control
mammary gland imaged with bright field microscopy. (B) MPE image of (A) showing the power of MPLSM to visualize collagenstructure. (C) H&E stain of mammary gland from a col1a1/col1a1 homozygote mouse. (D) MPE image of (C) showing increased
visualization of collagen density and structure and the irregular epithelial-stromal boundary, with epithelial cells invading into
the locally increased collagen stroma. (E-G) MPLSM/SHG image of live mammary ducts that are not fixed, sectioned, or stained
from wild type (E), heterozygous (F), and homozygous (G) col1a1 mice, showing increased collagen density in transgenic animals. (H) Z-stack of (F) spanning 340 μm (typical depth capability 350–440 μm) illustrating the structure of the glands and theirrelative location in the gland (see Additional file 4). Note: scale bar for MPE/SHG images equals 25 μm.
similar to observations from the Wnt-1 mouse tumors.
For instance, tracking the tumor-stromal boundaryaround individual tumors (Figure 5A–D) first revealed
regions of TACS-2 morphology, in which restraining collagen wrapped around a relatively smooth boundary (Figure 5A,B, and histogram: TACS-2). In regions of TACS-2,
cells had a mostly random alignment with a subset of cellsaligned in the same direction as the collagen (Figure
5A,B). The second region revealed a TACS-3 boundary
where collagen has been primarily aligned perpendicular
to the tumor (Figure 5C,D, and histogram: TACS-3). A reasonable hypothesis for this behavior is that collagen fiber
realignment occurs through morphogenesis and contractile events from cells at the tumor boundary, to organize
the matrix and to prepare for local invasion (see Figure 6
for TACS-2 to -3 transition). Evidence for this hypothesis
is seen by the collagen structure associated with regions ofinvading cells in the Wnt-1 tumors (Figure 4C), PyVT
tumors (Figure 5A–D; Figure 6), and MPLSM analysis ofinvasive regions in tumor histology sections (Additional
file 3). In all these cases local tumor invasion occurred
where collagen fibers are aligned radially from the tumor
in the direction of tumor cell invasion (see TACS-3 histogram in Figure 5, and the change in stromal alignment
correlated with the change to invasive cell morphology in
Figure 6). Additionally, some invading cells were in directcontact with fibers (Figure 5E,F; Figure 6C,D). In certain
cases, some matrix disorganization was also observable(see Figure 4C: c and Figure 5E) possibly indicating proteolytic cleavage of collagen [40,43].
To further test the hypothesis that tumor cells realign thecollagenous matrix to facilitate local invasion, we determined whether tumor cells could reorganize a random
collagen matrix. Analysis of tumor explants within type-I
collagen gels revealed radial collagen alignment at regionsof tumor cell invasion into previously randomly aligned
collagen gels (Figure 7), consistent with previous reports
analyzing fixed tumor explants in 3D reconstituted matrices [22], and supporting information about invasion
within intact live tissues (Figures 4, 5, 6). Following
polymerization, circular collagen gels (disks) displayed arandom orientation of collagen fibers (Figure 7A) without
an outside force to initiate structural reorganization.
Tumor explants cultured within such random collagengels contracted and reorganized these collagen gels, resulting in various local outcomes with collagen wrappedaround the explant at non-invading regions (Figure 7B).
In contrast, at regions of invasion into the gels, collagenwas reorganized to a radial alignment (Figure 7C) withdirect contact between collagen fibers and invading cells(Figure 7C,D). This demonstrates that tumor cells can dePage 8 of 16(page number not for citation purposes)

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FigureThetumor-stromal
4
interaction: representation of the three TACS in Wnt-1 mouse tumors
The tumor-stromal interaction: representation of the three TACS in Wnt-1 mouse tumors. (A-C) Micrographs
illustrating the identified TACS. A: TACS-1. MPE/SHG image of TACS-1. Namely, a region of dense collagen (a, and surfacemap) "above" a non-palpable tumor (b,c; yellow outline) that is indicative of the presence of a small tumor (also see Additional
file 5). The surface map quantifies the intensity of the fluorescent signal relative to x-y location, and clearly demonstrates anincreased collagen signal, and is representative of six Wnt-1 tumors and eight PyVT tumors (not shown) B: TACS-2. MPE/SHG
image of the second TACS indicated by the presence of straightened (taut) fibers characteristic of a larger Wnt-1 tumor. B: ac, MPE/SHG images of collagen fibers in Wnt-1 mice stretched around a relatively smooth tumor boundary (outlined with ayellow line in c) as demonstrated by the fact that majority of the fibers are parallel to the tumor boundary. B: histogram, theangle of collagen fibers relative to a line tangential to the tumor boundary was measured for 86 regions in six independent
tumors, and graphed as a frequency distribution resulting in a distribution of fibers around 0°. C: TACS-3. The third TACS:aligned collagen fibers at regions of cell invasion in Wnt-1 mice. C: a-c, he irregular tumor boundary associated with local invasion is outlined in yellow (in a) and connected to fibers that are primarily distributed normal to the initial tumor boundary, represented by a frequency distribution around 90° relative to the tumor boundary. C: histogram, the angle of collagen fibers
relative to a line tangential to the tumor boundary was measured for 71 regions in six independent tumors, and graphed as afrequency distribution resulting in a distribution of fibers near 90°. Scale bars equal 25 μm.

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Figureof5TACS-2 and -3 in the more aggressive PyVT mouse tumor model
Study
Study of TACS-2 and -3 in the more aggressive PyVT mouse tumor model. (A) TACS-2, with an enlarged cutout
region (B) shown at higher brightness and contrast levels and a rough demarcation of the tumor-stromal boundary (red line; s= stroma; t = tumor) to further indicate the wrapping of the collagen parallel to the tumor boundary (distribution near 0°, see
top right TACS-2 histogram of 106 regions from eight independent tumors). (C) TACS-3, with an enlarged cutout region (D)shown at higher brightness and contrast levels and a rough demarcation of the tumor-stromal boundary (red line; s = stroma;t = tumor). Note that although some cells have moved past this boundary (examples = x) into the fibers, the boundary serves
as a general representation of irregular invasive region into radially aligned collagen fibers (frequency distribution around 90°,see middle right TACS-3 histogram of 109 regions from eight independent tumors). Furthermore, analysis of TACS-3 regions(E) at higher magnification (F) show endogenous cells associated with fibers at the tumor-stromal boundary and within thetumor. * Indicates examples of fibers interdigitated with the invasive tumor boundary and in contact with the invading tumor
cells (red arrows). Scale bar for MPE/SHG images A, C, E, and F equals 25 μm, and 10 μm for B and D.

novo reorganize a random collagen matrix to facilitate
invasion, supporting the hypothesis that matrix reorganization at the tumor interface facilitates local invasion in

vivo. Hence, our results suggest that in order to facilitate
local invasion, cells at the tumor boundary contract andalign collagen fibers, perhaps with the assistance of prote-

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Figure 6– radially aligned collagen fibers associated with invasion
TACS-3TACS-3 – radially aligned collagen fibers associated with invasion. Combined MPE/SHG imaging of live intact PyVT
mammary tumors indicates that (A) non-invading regions (*) possess taut collagen (TACS-2; white arrowhead) wrappedaround the tumor, while regions of invasion (x) are linked to aligned collagen, with cells invading along radially aligned collagenfibers (TACS-3; black arrowheads). Examination of invasion at the tumor-stromal interface at higher magnification (B and C)
clearly reveals the collagen alignment at specific regions of invasion (B) with tumor cells (*) in between, and in association with,aligned collagen fibers (C). (D) Example of an individual tumor cell attached to a collagen fiber leading away from the primarytumor. Additionally, tumor cells that have invaded across the tumor-stromal boundary can be visualized by separating the MPE
and SHG signals or imaging with FLIM. (E) Combined MPE/SHG image of a TACS-3 region facilitating local invasion. The tumorstromal boundary is not well preserved at this stage, but is roughly outlined in red. Examples of regions of cells that haveinvaded past the boundary are marked with an asterisk (t = the primary tumor). The red arrowhead indicates cells near thetumor boundary that are migrating along aligned collagen fibers away from the primary tumor. (F) MPE and SHG signal separation of the image shown in E. MPE signal is represented in red pseudo-color while SHG is shown in green pseudo-color. Notethe interdigitation of aligned collagen fibers (green) into the tumor, with individual, or lines of, cells (* in E) migrating away fromthe tumor on collagen fibers. (G) FLIM micrograph of invading cells at the TACS-3 region shown in E and F. Collagen (blue; nofluorescence lifetime) can be distinguished from cells (green to yellow), confirming the presence of invading cells at the tumorstromal boundary and cells that have migrated past the boundary in association with collagen. The color bar in G ranges represents the weighted mean ranging from 100 ps (blue) to 1 ns (red). Scale bars equal 25 μm in A, E, F, and G; 10 μm in B, C, andD.

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Figure 7matrix reorganization by tumor cells facilitates local invasion of tumor cells
Collagen
Collagen matrix reorganization by tumor cells facilitates local invasion of tumor cells. Combined MPE/SHG imaging of tumor explants cultured within 3D collagen gels for eight hours demonstrates that tumor cells reorganize a previously
random matrix to facilitate invasion. (A) Region of 3D collagen gel remote from the tumor demonstrating the random orientation of collagen present within 3D collagen gels in a region that has not been reorganized by a specific outside force. This random organization is specifically altered by cells from tumor explants as the cells contract and reorganize the collagen matrix.
Similar to data in live intact tissues, non-invading regions show collagen pulled in near the explant but wrapped around thetumor boundary (B), while at regions of tumor cell invasion into the collagen gel, collagen has been radially aligned by thetumor cells (white arrow heads C) with cells (*) in direct contact with the collagen matrix (white arrowheads; D). Therefore,
at regions of tumor cell invasion, collagen has been reorganized to a radial alignment from a random orientation, indicating a
structural realignment of collagen fibers facilitates local invasion. Scale bar equals 25 μm.

olytic cleavage to facilitate matrix reorganization, andthen invade along aligned collagen structure to expand
the tumor and later metastasize.

Discussion
Intrinsic fluorescence detection with multiphoton excitation in combination with SHG facilitates three-dimensional, high resolution, imaging in unfixed, unsectioned,

unstained mammary tissues. This imaging provides information commonly obtained with classical histology andEM without the need for complex and destructive sample
preparation, and with additional structural information
in four dimensions. With MPE/SHG, mammary gland tissue could be clearly imaged at depths of 440 nm, andchanges in collagen density could be reliably detected.
Furthermore, three-dimensional imaging of tumors inPage 12 of 16(page number not for citation purposes)

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situ revealed three now defined TACS, which provide
standard hallmarks to locate and characterize tumors:
TACS-1, the presence of dense collagen, indicated byincreased signal intensity at a region around the tumor asa standard hallmark for locating small tumor regions;
TACS-2, the presence of taut (straightened) collagen fibers
stretched around the tumor, indicating growth leading toincreased tumor volume; and TACS-3, the identificationof radially aligned collagen fibers facilitating invasion,
which may be indicative of the invasive and metastatic
growth potential of a tumor. Together these signatures
may serve a mechanism to help identify and characterize
breast tumors in experimental animal models as well as
human cancers and fresh tumor biopsies.
The breast epithelial cell-ECM interaction is responsiblefor influencing cell polarity, proliferation, differentiation,adhesion, and migration [44,45] and type I collagen is an
important regulator of mammary ductal formation during
development [4]. Analysis of normal mammary glands
reveals collagen fibers wrapping around, as well as radiating away from, the duct (Figure 2A: b). This organizationis remarkably consistent with the observation that in fixedwhole mounts of developing mammary gland, analyzed
with multiphoton microscopy, collagen fibers are "pulledin" perpendicular to the terminal end bud [46], similar towhat we observe for radially aligned collagen fibers neartumors (TACS-3). Combined, these morphologies provide insight into the structure-function relationship in the
mammary gland and imply that collagen may provide
directional cues during development that also influencechanges in the normal mammary gland. For instance, thecrimped (wavy) collagen structure (i.e. Figure 2A: a,b,e,g)
is consistent with numerous reports of crimped collagenfibers in connective tissue that allow normal tissue deformation with a strain-stiffening behavior [47,48]. Thisbehavior may hold true for the mammary gland as well,allowing for tissue deformation and normal ductal
growth and involution without over constraining the system, yet providing adequate levels of tensile resistance tocontracting cells and resisting large deformations that candamage the tissue. The less numerous taut fibers may
serve a different purpose. They may act as locally constraining structures at the single cell level and may act tointerconnect various ducts in the tissue together and to thenipple structure (Figure 2A: f), which may transmit
mechanical signals to the ducts during activities such as
nursing to elicit mechanotransductive signaling related tolactation. Furthermore, such mechanical signals acting
directly on epithelial cells or transmitting stress across thebasement membrane would be amplified by increased
breast tissue density. Hence, increased breast tissue density in vivo may promote carcinoma formation byincreased mechanical signaling events in dense tissue,

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consistent with in vitro work showing that increased
matrix density alters breast epithelial cell signaling [49].
The importance of matrix composition and morphology
around the mammary epithelium is illustrated by studies
showing that misregulated stromal-epithelial interactions
can promote tumorigenesis [6-8] and the fact that breast
carcinomas often exhibit desmoplasia (excessive collagensurrounding an invasive tumor [50]). Moreover, cancer
cells can locally invade across basement membrane andcollagenous stroma to spread into neighboring ECM environments, where they can migrate further to enter lymphatic and blood vessels, resulting in metastatic growth in
distant tissues [40,51]. Therefore, understanding themechanisms of invasion in vivo is of great importance.
Yet, to our knowledge, no study has visualized local invasion in endogenous tumors in vivo in relation to stromalorganization. Consequently, it is noteworthy that we
observe alignment of collagen fibers, and association ofindividual cells with those fibers at regions of local invasion in live tissue (TACS-3), which is similar to observations of individual cell migration along collagen fibers ina xenograft model [9], and confirms and expands upon invitro studies in 3D matrices that have identified collagen
reorganization (alignment) at the front of invading cells[22,40,41]. Moreover, the concept of alignment-facilitated invasion appears to be of significance in collective
cell migration (e.g. tubulogenesis in the mammary gland;[41]) as collagen alignment is noted at the terminal end
bud during invasion of the mammary ductal tree [46].
Thus, collagen alignment may facilitate motility andmigration during normal development, while tumor invasion may resemble misregulated developmental processes.

Conclusion
The data presented indicate that tumor cells often localize
near dense collagen or promote a desmoplastic response
and contract and localize collagen, followed by tumor
growth and expansion (stretching) of the collagen matrix
leading to matrix reorganization (possibly assisted by proteolytic cleavage [40,43] to release collagen fibers) to help
facilitate local invasion. This matrix reorganization would
require enhanced contractility and motility of the tumor
cells, which may explain the increased presence of Rho
and ROCK, in invasive cancers ([52], and references
therein). Although a number of mechanisms, such as
growth factor and integrin signaling and protease secretion and activity, are associated with invasion and metastasis, it seems likely that GTPase-regulated motility events
are also involved these processes. Hence, the mechanisms
behind local invasion may include matrix reorganization
through GTPase-mediated tumor cell contractility (P.P.P.
and P.J.K unpublished observations), leading to analigned matrix that facilitates local invasion.

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Competing interests

Additional File 4

The authors declare that they have no competing interest.
However, portions of the technologies presented in the
manuscript are patent pending. The authors have no interest, arrangement, or affiliation that could be perceived asa conflict of interest in the context of this manuscript.

Movie file showing MPE/SHG imaging of collagen surrounding the mammary duct at >440 μm into the live tissue.
Click here for file
[http://www.biomedcentral.com/content/supplementary/17417015-4-38-S4.avi]

Authors' contributions

Additional File 5

PPP conducted all MPLSM, SHG, SEM, and histology
experiments, managed mouse colonies, performed 3D
cell culture experiments, analyzed the imaging data, andprepared the manuscript and figures. DRI assisted withmice and performed 3D culture experiments. JMC performed TEM imaging of mouse glands. KWE and JGWassisted with specific technical aspects of nonlinear imaging and data analysis as well as project coordination. PJK
participated in the design and coordination of the project
and assisted with data analysis. PPP, KWE, JGW, and PJK
cooperatively designed the project and discussed datainterpretation and analysis. All authors participated in
critical editing of the manuscript and read and approved
the final manuscript.

Movie showing MPE/SHG imaging of TACS-1.
Click here for file
[http://www.biomedcentral.com/content/supplementary/17417015-4-38-S5.avi]

Additional material
Additional File 1
MPM/SHG imaging of collagen and endogenous fluorescence in connective tissue and reconstituted 3D matrix. (A) SHG image of type I collagenin mouse Achilles tendon. (B) MPM/SHG image of cellular endogenous
fluorescence and collagen in skin. (C) Collagen and NADP(H) detection
with combined MPM/SHG imaging of mouse pectoral muscle. (D) SHG
imaging of type I collagen in a reconstituted three-dimensional gel.
Click here for file
[http://www.biomedcentral.com/content/supplementary/17417015-4-38-S1.pdf]

Acknowledgements
The authors thank Drs Caroline Alexander and Amy Moser for providingcolony founder mice, Curtis Rueden for assistance with VisBio, and theMaterials Science Center and BBPIC for EM resources. This work was supported by grants from the DOD-CDMRP/BCRP: W81XWH-04-1-0428 to
PPP, the S.G. Komen Foundation: BCTR02-1841 and NIH: CA076537 to
P.J.K. and NIH NIBIB: R01-EB000184 to J.G.W and K.W.E.

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Additional File 2
Col1a1tmJae mice possess collagen dense mammary tissue. Histology ofhomozygous col1a1 mice (A) showing increased collagen surrounding the
mammary duct as detected with H&E, trichrome, and picrosirius red(Picro) staining. (B) Increased collagen is also present in heterozygous
col1a1 mice as detected with H&E and picrosirius red staining (PS).
Click here for file
[http://www.biomedcentral.com/content/supplementary/17417015-4-38-S2.pdf]

Additional File 3
MPLSM examination of TACS-2 and TACS-3 in H&E sections of PyVT
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[http://www.biomedcentral.com/content/supplementary/17417015-4-38-S3.pdf]

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