Located in Barnard Hall next to the Strand Union Building, the Center for Biofilm Engineering comprises more than 20,000 square feet, and includes offices and conference rooms for faculty, staff, and students; a computer lab; and 15 fully equipped research laboratories and at least 9 additional directly affiliated laboratory spaces. CBE Core Facility labs include an analytical instrument lab, a microbiology lab with media preparation area and autoclaves, microscope facilities, as well as an isolated radioactive isotope lab with liquid scintillation counter. See below for a comprehensive list of shared equipment available.

Core Facilities

Analytical and Molecular Core

The analytical core lab is a dedicated space with instruments that are maintained by the CBE Technical Operations Manager.  Users including students, staff and faculty are trained to prepare their samples and standards, run instruments, analyze data and troubleshoot data analysis and methods by the TOM. There are three gas chromatographs with detectors including Mass Spectrometer, Thermal Conductivity and tandem thermal conductivity/Flame Ionization for analysis of permanent gases, ethylene/acetylene or fatty acid methyl esters on the MS detector. Liquid chromatography capabilities are extensive with two instruments that can be configured for various analysis including amino acids, organic acids, alcohols, carbohydrates, cannabinoids, industrial compounds of interest and photosynthetic pigments. The more basic system includes tandem Variable Wavelength Detector (UV) and Refractive Index Detector with a high-pressure quaternary pump.  The more advanced HPLC has a temperature controlled, programmable autosampler for performing pre-column derivatizations in the sample needle, for detection with the highly sensitive and tunable multi-channel Fluorescence Detector. There is a dedicated Anion Ion Chromatography system and Total Carbon Analyzer that is configured for either non-purgeable organic carbon or dissolved inorganic carbon measurements.  Spectrophotometers in the core are capable of visible and fluorescence measurements in vessels including cuvettes, test tubes and microwell plates.  The plate reader can also measure luminescence as perform kinetic time scans. For small measurements a microbalance is available as well as a micro pH meter that can measure pH in as little as 10uL.

The primary molecular core lab includes advanced instrumentation for nucleic acid, extraction and detection, including an Illumina MiSeq Sequencing System as described in further detail below. The molecular core also includes an Agilent 2100 Bioanalyzer, MP Biomedical FastPrep 24 beadbeater and Oxford Nanopore Technologies MinION real-time sequencing device.  A molecular satellite station includes two thermocyclers, a gel running and imaging station, and spectrophotometers for nucleic acid quantification. Contact: Kristen Brileya

Microscope Core

The microscopy and chemical imaging facilities are coordinated by the Microscopy Facilities Manager who maintains the equipment and trains and assists research staff and students in capturing images of in situ biofilms via optical microscopy, fluorescent and Raman confocal microscopy. The microscopy facilities include four separate laboratories—the Optical Microscopy Lab, the Confocal Microscopy Lab, the Chemical Imaging Lab, and the Microscope Resource Room and Digital Imaging Lab—which are detailed below.

The Optical Microscopy Lab houses a newly acquired Leica Thunder widefield microscope optimally configured for allow real-time, high-sensitivity, high-throughput imaging. This microscope is equipped with a cMOS Camera, a Tunable multiline LED Light Source, an ultraprecise, triggerable stage, has complete Hardware Synchronization and is fully enclosed to maintain temperatures between ambient 5-50C (± 0.1°C), humidity (specifically designed to prevent condensation), and CO2/Air or hypoxic/hyperoxia conditions, and real time computational removal of out of focus background and increased imaging depth of thicker specimens. The acquisition speed is nearly five times faster than a conventional Confocal Scanning Laser Microscope (CSLM). Experiments that would normally take hours to visualize on a CSLM can be accomplished in minutes by the widefield Thunder microscope. For example, to image 96 wells, one position per well, 10 z-planes per well, and three channels per well would take only 3.5 min on the proposed THUNDER imager system. This system is ideally configured for improved live cell viability for longer term imaging studies due to advanced ‘under microscope’ incubation capabilities and reduced phototoxicity and photobleaching by limiting light doses.

An upright, Leica DM6 fluorescent microscope that is a fully automated with constant color temperature and fully automated transmitted light- and fluorescence axis, with motorized Z-focus.

A Nikon Eclipse E-800 research microscopes which are used for transmitted light and epi-fluorescent imaging. Both microscopes are equipped with Photometrics MYO cooled CCD cameras and use Universal Imaging Corporation’s MetaVue software (v 7.4.6) for digital image acquisition. We have a large collection of fluorescence filter cubes for the Nikons, including those optimized for the following fluorescent stains: FITC (gfp), TRITC (propidium iodide), DAPI, CTC, ELF-97, CY5, cfp, and we also have a B2E cube. Both Nikons are equipped with Nomarski/DIC, and we have a 100x oil phase contrast objective and condenser especially for use with imaging spores.

A Leica LMD6 Laser Microdissection System equipped with a color camera, fluorescence filter cubes (FITC, TRITC, DAPI), and a UV laser for sample dissection.

A GAN210 Optical Coherence Tomography (OCT) imaging system. OCT is a high resolution, non-contact, non-invasive, and non-fluorescent based technique that is well suited for imaging thick specimens. The OCT light source centered around 930 nm with a bandwidth >100 nm and has a scan rate of up to 36 kHz with an axial field of view of 2.9 mm / 2.2 mm. Depending on the scan objective the field of view (FOV) and resolution can be adjusted and vary between a larger FOV of 16x16 mm² at 12 µm resolution, and a FOV of 10x10 mm² with a higher resolution of 8 µm.

Additionally, within the Optical Microscopy Lab is a Leica M 205 FA computer-controlled stereomicroscope and a Leica DFC3000G fluorescence camera. This stereoscope can be used to image samples using fluorescence, brightfield with or without polarization or Rotterman contrast, and reflected white light. The software will also allow a z-stack of images to be collected and recombined using simple deconvolution. Other equipment in the Optical Microscopy Lab includes a Nikon SMZ-1500 barrel zoom stereomicroscope equipped with a color camera, a Leica cryostat, and a dry ice maker.

The Confocal Microscopy Labcontains two Leica SP5 Confocal Scanning Laser Microscopes (CSLMs).  One is an inverted confocal microscope with 405, 488, 561 and 633 nm laser excitation lines. It is equipped with a tandem scanner, so it can be switched from standard scanning mode to operate in Resonant Scanner mode, which enables scanning at exceptionally high frequencies for fluorescent imaging. This faster scanning is necessary for most live cell imaging (note: “live cell imaging” doesn’t generally refer to imaging bacterial cells, but rather mammalian cells and processes). This inverted SP5 also includes a heated stage with an environmental control chamber (i.e. it can be used to provide an enclosed CO2 atmosphere), and a motorized stage with Mark-and-Find and image tiling capabilities.

The second SP5 is an upright confocal microscope, also with 405, 488, 561 and 633 nm lasers, a motorized stage, Mark-and-Find, and tiling capabilities. This upright has a removable heated chamber that encloses the entire microscope, so that larger, incubated flow cell systems can be accommodated over long periods of time. This enables high-resolution time-lapse monitoring of biofilm development, treatment and detachment phenomena. Additionally, this microscope is equipped with Fluorescence Lifetime Imaging (FLIM) capability, which is also referred to as Single Molecule Detection.

The CSLM is capable of imaging biofilms on opaque surfaces, so a wide variety of materials can be used in the experimental flow cells. As biofilm formation proceeds in an experiment, representative areas of the colonized surface are scanned with the use of the automatic stage. Digital data is collected from sequential scans, and stored data can be viewed in the x, y, z coordinates to yield a 3-dimensional image of the biofilm architecture. Quantitative and qualitative information about biofilm architecture can be retrieved easily from examination of CSLM data, in both the x-y and x-z planes, and the existence or absence of structural features, such as microcolonies and water channels, can be determined.

TheChemical Imaging Laboratorycontains two Raman Microscopes. One is a Horiba LabRam HR Evolution NIR that is dedicated to studying the molecular composition of a sample. This is a fully integrated high resolution Raman microscope for confocal Raman analysis, optimized for the visible to IR range (400nm-2500nm) microscope. Includes confocal Raman microscope with an automated xyz-stage with fast-mapping capabilities, transfer optics, stigmatic spectrometer equipped with two gratings (600 and 1800 l/mm gratings), multichannel air-cooled CCD detector, and computer package with the latest version of the LabSpec6 software and the KnowItAll Raman spectra library, Horiba edition. It is equipped with 532nm 100mW laser, HeNe 633nm laser, 785nm 90mW laser, and 10x, 50x, 100x, 20xLWD and 50xLWD objectives. The second Confocal Raman Microscope is dedicated to studying the metabolic activity of single microbial cells as well as a cell’s health/disease state. This is a modified Horiba LabRam Evolution Confocal Raman microscope and is equipped with a 500mW 532nm laser, a 100mW 785nm laser, ultra-low frequency filters for stokes and anti-stokes measurements, an air-cooled open electrode CCD, a deep-cooled back-illuminated EMCCD multichannel detector, 300 and 1800 gr/mm grating, an automated xyz-stage with fast-mapping capabilities, filter sets for FITC, CY3, DAPI, and Cy5, 10x, 50x, 63x (long working distance), and 100x objectives, and a CaptuR laser trapping (1,500mW, 1,064nm) system (=optical tweezer). The computer workstation is equipped with the latest version of the LabSpec6 software with an integrated multivariate analysis module, a particle finder module, as well as a KnowItAll Raman spectra library, HORIBA Edition.

The Microscope Resource Room / Digital ImagingLab is where CBE researchers examine and reconstruct the stacks of image data they have collected using our image analysis software. For quantitative analysis, such as intensity or particle-size measurements, we use Universal Imaging Corporation’s MetaMorph software. We use Bitplane’s Imaris software for computer-intensive data analysis like particle tracking and for qualitative analysis—for example, putting together a stack of 200 red and green flat images to get a 3-dimensional image of a biofilm microcolony that can be rotated in space and examined from every angle. The lab consists of three dedicated computers, a server for storing large files, CD and DVD burners and readers, and a color printer. In addition to providing CBE students, staff, and researchers with an imaging workplace, the resource room gives us a place to hold group tutorials and WebEx group software training sessions.

Contact for Microscopy Core:Heidi Smith

Specialized CBE Laboratories

Ecology/Physiology Laboratory

The Ecology/Physiology Laboratory led by Dr. Matthew Fields has general microbiology equipment, anaerobic gassing stations in two lab spaces, Ultra-Centrifuge, Anaerobic Chamber, biofilm reactors, protein and DNA electrophoresis, Qubit fluorometer, two Eppendorf Mastercylcers, and a microcapillary gas chromatograph with dual TCDs. The lab has light-cycle controlled photo-incubators as well as photo-bioreactors for the cultivation of algae and diatoms.

This laboratory houses an Illumina MiSeq Sequencing System in its shared molecular core area. The MiSeq desktop sequencer allows the user to access more focused applications such as targeted gene sequencing, metagenomics, small genome sequencing, targeted gene expression, amplicon sequencing, and HLA typing. This system enables up to 15 Gb of output with 25 M sequencing reads and 2x300 bp read lengths by utilizing Sequencing by Synthesis (SBS) Technology. A fluorescently labeled reversible terminator is imaged as each dNTP is added, and then cleaved to allow incorporation of the next base. Since all four reversible terminator-bound dNTPs are present during each sequencing cycle, natural competition minimizes incorporation bias. The end result is true base-by-base sequencing that enables the industry's most accurate data for a broad range of applications. The method virtually eliminates errors and missed calls associated with strings of repeated nucleotides (homopolymers). Contact:Sara Altenburg

Medical Biofilm Laboratory

The Medical Biofilm Laboratory (MBL) has earned a reputation for being a university lab that focuses on industrially relevant medical research in the area of health care as it relates to biofilms. Dr. Garth James (PhD, microbiology), Randy Hiebert (MS, chemical engineering), and Dr. Elinor Pulcini (PhD, microbiology) have been the innovative leaders and managers of this respected, flexible, and adaptable lab group. The MBL team also includes a full-time Associate Research Professor, a Research Professional, a Research Associate, and two Undergraduate Research Assistants.

Currently, eighteen companies, including CBE Industrial Associates, sponsor MBL projects. These projects include evaluating antimicrobial wound treatments and dressings, prevention of biofilm formation on medical devices, evaluation of biofilm formation in endoscopes, testing endodontic irrigants, evaluating virus transfer from surfaces, and testing biofilm prevention and removal agents. The MBL is a prime example of integration at the CBE, bringing together applied biomedical science, industrial interaction, and student educational opportunities. Contact: Garth James

Standardized Biofilm Methods Laboratory

The Standardized Biofilm Methods Laboratory (SBML) was designed to meet research and industry needs for standard analytical methods to evaluate innovative biofilm control technologies. SBML staff and students develop, validate, and publish quantitative methods for growing, treating, sampling, and analyzing biofilm bacteria. The SBML members work with international standard setting organizations (ASTM International, ISO) on the approval of biofilm methods by the standard setting community. Under a contract with the U.S. Environmental Protection Agency (EPA), the SBML provides statistical services relevant to the EPA's Office of Pesticide Programs Microbiology Laboratory Branch to assess the performance of antimicrobial test methods—including those for biofilm bacteria. The SBML received funding from the Burroughs Wellcome Foundation to develop a method for assessing the prevention of biofilm on surface modified urinary catheters that was approved in 2021 as ASTM Standard Test Method E3321.  In addition, they conduct applied and fundamental research experiments and develop testing protocols for product specific applications. Methods include: design of reactor systems to simulate industrial/medical systems; growing biofilm and quantifying microbial abundances and activity; testing the efficacy of chemical constituents against biofilms; and microscopy and image analysis of biofilms. SBML staff offer customized biofilm methods training workshops for CBE students, collaborators, and industry clients. Contact: Darla Goeres

Microbial Ecology and Biogeochemistry Laboratory

Research in the Microbial Ecology and Biogeochemistry Laboratory (www.foremanresearch.com) lies at the intersection of microbial ecology and engineering and uses a combination of field and laboratory studies, as well as approaches ranging from the single-cell to the community level. Staff in this lab are interested in understanding how the environment controls the composition of microbial communities and how, in turn, those microbes regulate ecosystem processes such as nutrient and organic matter cycling. Ongoing research examines carbon flux through microbial communities, with the long-term goal of improving predictions of carbon fate (metabolism to CO2, sequestration into biomass, long-term storage in ice) in the context of a changing environment. Additionally, the lab is interested in physiological adaptations to life in extreme environments, biofilms in space, microbial biosurfactants, genomics and spectral detection of organic compounds. Contact: Christine Foreman

Environmental Sensing Laboratory

Sensors are essential for understanding and predicting environmental changes by measuring biological, chemical, and physical properties. The Warnat laboratory develops microfabricated sensor systems that allow in situ measurements with a high spatial and temporal resolution in various harsh environments such as water systems, snow and ice, soil, or maple syrup lines. Sensors can be integrated into microfluidic environments that allow measurements of ultra-small volumes and simultaneous visualization of biological processes on the sensor surfaces using high-resolution microscopy. Ongoing research examines in collaboration with CBE colleagues how fabricated sensor systems can be integrated into various biofilm-forming environments to detect initial biofilm attachment and provide an electrical feedback signal for potential biofilm mitigation strategies. Contact: Stephan Warnat

Microfluidics Laboratory

Dr. Connie Chang runs a soft materials and microfluidics laboratory to study microbes (bacteria, biofilms, and viruses). Dr. Chang is applying drop-based microfluidics—the creation and manipulation of picoliter-sized drops of fluid—for high-throughput screening and assaying in biology. Her lab is developing novel tools for quantifying the behavior of individuals and how they can collectively contribute to large-scale population dynamics. Ongoing projects within her group include the screening of persister and dormant bacteria cells in biofilms and the study of influenza evolution and population dynamics.

Dr. Chang has shared laboratory space in the CBE and an individual laboratory space in the Chemistry and Biochemistry Building (CBB) at MSU. The laboratory spaces include common space for equipment, chemical storage, freezers and reagents. The lab is outfitted with a qPCR machine and also includes a dedicated a room for epifluorescence microscopy and a custom built microscope stand (200 square feet). The lab contains all the equipment and instrumentation necessary for fabrication of new devices, microfluidics handling, PCR, and cell culture. Contact:Connie Chang

Bioprocess and Biofilm Technology Laboratory

Dr. Gerlach oversees the Bioprocess and Biofilm Technology Laboratory (BBTL), which is a set of laboratories focusing on the development of engineering applications, relevant for industry, the environment and medicine. The BBTL develops and improves engineering processes based on the use of traditional chemical, biological and mechanical process-schemes through combination with biofilm- and biomineralization-specific aspects. Work in the BBTL cuts across all domains of life with current foci on fungi, algae, and -of course- bacteria and archaea. The BBTL was essential in the development and commercialization of a biocement-based well-sealing technology (BioSqueeze®) and is currently focusing on the development of biocement-based infrastructure materials. Algal biofuel and bioproduct generation are additional research and development topics with a current focus on high pH/high alkalinity adapted extremophiles as well as the capture of carbon dioxide directly from the air. The BBTL facilitates fundamental and applied research, and has specialized equipment available that includes small and large-scale, high pressure and high temperature bio- and biofilm-reactors and incubators capable of supporting (photo)autotrophic and heterotrophic growth experimentation, porous media micromodels, flowcells, etc. suited for the cultivation of biofilms and microorganisms, Gas Chromatographs with Mass Spectrometric (GC-MS), electron capture (ECD) and flame ionization detectors (FID), an elemental analyzer (EA), a Thermal Gravimetric Analyzer (TGA), a Raman microspectroscopy instrument, a Fourier-Transform Infrared (FTIR) spectrometer, an ion chromatograph, an automated titrator, fluorescence and absorbance plate readers, as well as all necessary standard chemical analysis and cultivation capabilities necessary for biofilms and microbe cultivation, including capabilities for the cultivation of microaerophilic or anaerobic microbes (modified Hungate setup and anaerobic glovebag). The use of advanced molecular biology techniques including next generation sequencing for community analyses, metagenomics and transcriptomics are applied routinely in combination with next generation physiology techniques. Contact: Robin Gerlach

Microsensor Laboratory

The Microsensor Laboratory provides the capability of measuring microscale chemical and physical parameters within biofilms, microbial mats and other compatible environments. The Microsensor Laboratory has the capability to measure spatial concentration profiles using sensors for oxygen, pH, hydrogen sulfide, nitrous oxide and some custom-made electrodes. All electrodes are used in conjunction with computer-controlled micromanipulators for depth profiling. A Leica stereoscope is used to visualize the sensors while positioning them on the biofilm surface. The laboratory has experience with diverse microsensor applications including biofilms in wastewater, catheters and hollow fiber membrane systems in addition to algal and fungal biofilms. Contact:Kristen Brileya

Environmental Microbiology Lab

​The research activities of the Environmental Microbiology Lab headed by Dr. Roland Hatzenpichler focuses on microbial ecophysiology, the study of the physiology of microorganisms with respect to their habitat. We are interested in how the activity of the “uncultured majority” – the large number of microbes that evades cultivation under laboratory conditions – impacts humans and the environment on a micron to global scale. We are convinced that only by gaining an understanding of microbes directly in their habitats researchers will be able to elucidate the mechanisms of microbial interactions with the biotic and abiotic world. To accomplish these goals, we apply an integrative approach that bridges the two extremes of the microbial scale bar: the individual cell and the whole community.​

Very broadly, the research questions our lab addresses are: (1) who is doing what (linking phylogenetic identity and physiological function), (2) what are the abiotic and biotic factors controlling microbial in situ activity, (3) how does this activity affect the environment and ultimately humans, (4) what are the limits to metabolism in terms of energy, space, and time, and (5) how can we discover novel structures and functions within uncultured microbial lineages?​​

Our approach to these problems is inherently multi-disciplinary and multi-scaled. In order to address previously unrecognized physiologies and cellular interactions of uncultured microbes, we employ a unique combination of metagenomics (as hypotheses generator), high-through-put metabolic screening via substrate analog probing (to identify geochemical and biotic parameters driving ecology), and single cell resolved stable isotope probing via Raman microspectroscopy or nano-scale secondary ion mass spectrometry (to identify specific growth-sustaining substrates). These culture-independent approaches are complemented by mesocosm experiments run under close to in situ conditions and targeted cultivation efforts. Because, together, these approaches target the whole microbiome as well as the individual cell we typically do not depend on samples enriched in a target population, as is often necessitated in ecological studies. Our main study sites are sediments from a variety of geothermal, deep-sea, and coastal habitats. Contact: Roland Hatzenpichler

CBE Computer Facilities

The CBE maintains several dedicated computational and data storage computer systems including 6 high performance data and image analysis workstations and servers in addition a 143TB allocation on centrally maintained high throughput backed up high availability storage systems. The center provides personal workstations for staff and graduate students that are connected to the MSU computer network. A student computer laboratory offers nine state-of-the-art PCs along with scanning and printing services. Additionally, CBE staff and students have access to the centrally maintained computational cluster for data manipulation, analysis, and mathematical modeling. This cluster consists of 77 nodes with a total of 1300 hyper-threaded cores and 22 teraflops of computing power.

OTHER Montana State University facilities available for collaborative research

Montana Nanotechnology (MONT) Facility

he MONT facility was formed from a $3 million NSF grant awarded to MSU in September of 2015. This collaborative facility includes the Montana Microfabrication Facility (MMF), the Imaging and Chemical Analysis Lab (ICAL), the CBE, the MSU Mass Spectrometry facility, and the Center for Bio-Inspired Nanomaterials. MONT provides researchers from academia, government and companies large and small with access to university facilities with leading-edge fabrication and characterization tools, instrumentation and expertise within all disciplines of nanoscale science, engineering and technology. Contact:David Dickensheets

MSU Nuclear Magnetic Resonance (NMR) Facility

A state-of-the-art NMR facility is available on campus on a recharge basis for research projects. This facility is a 5-minute walk from the College of Engineering and CBE laboratories. All the instruments in the facility are Bruker Avance instruments. The facility houses 300, 500 and 600 MHz NMR instruments for high resolution spectroscopy analysis. Contact:Valerie Copie

MSU Magnetic Resonance Microscopy (MRM) Facility

A state-of-the-art MRM facility is available on a recharge basis for research projects. This facility is located in the College of Engineering in the same building as the Center for Biofilm Engineering. Instruments in the facility are Bruker Avance III 250 MHz standard/wide bore and 300 MHz wide/super-wide bore spectrometers with Microimaging probes for each configuration. The facility provides measurements of NMR relaxation and diffusion to characterize molecular dynamics, e.g. for microscale EPS gel structure characterization and mesoscale MR imaging of heterogeneity in molecular dynamics and bulk scale transport phenomena and fluid dynamics. The imaging systems are capable of generating MRI and transport data with spatial resolution on the order of 10 μm in a sample space up to 6 cm diameter in opaque samples. Contacts:Sarah Codd and Joe Seymour

MSU ICAL Laboratory

The Imaging and Chemical Analysis Laboratory (ICAL) in the Physics Department at Montana State University is located on the 3rd floor of Barnard Hall, adjacent to the Center for Biofilm Engineering. ICAL is a user-oriented facility that supports basic and applied research and education in all science and engineering disciplines at MSU. The laboratory provides access to state-of-the-art equipment, professional expertise, and individual training for government and academic institutions and the private sector. Laboratory instrumentation is dedicated to the characterization of materials via high-resolution imaging and spectroscopy. ICAL promotes interdisciplinary collaboration between the research, educational and industrial fields,education, and industry; and the strengthening of existing cooperation between the physical, biological, and engineering sciences by providing critically needed analytical facilities. These facilities are open to academic researchers.

     ICAL currently contains eleven complementary microanalytical systems:

  • Atomic Force Microscope (AFM)
  • Field Emission Scanning Electron Microscope (FE SEM)
  • Scanning Electron Microscope (SEM)
  • Small-Spot X-ray Photoelectron Spectrometer (XPS)
  • Time-of-Flight Secondary Ion Mass Spectrometer (ToF-SIMS)
  • X-Ray Powder Diffraction Spectrometer (XRD)
  • Scanning Auger Electron Microprobe (AUGER)
  • Epifluorescence Optical Microscope
  • Critical Point Drying
  • Video Contact Angle System

For more information on each system, see the ICAL web site at: http://www.physics.montana.edu/ical/