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Porous polycarbonate membrane

Manufactured by Merck Group
Sourced in United States

The porous polycarbonate membrane is a lab equipment product that serves as a filtration material. It is made of polycarbonate and features a porous structure, allowing for the effective separation and filtration of various substances during laboratory processes.

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6 protocols using porous polycarbonate membrane

1

Cell Migration and Invasion Assay

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A total of 4×104 cells in 200 μl serum-free medium were added to the upper chamber of 24-well culture inserts with a porous polycarbonate membrane that was (8.0 μm, Millipore, Billerica, MA, USA) pre-coated (for invasion) with 30 μl Matrigel (BD Biosciences, San Jose, CA, USA) or not pre-coated (for migration) for 3 hours, and complete media was placed in the lower chamber. The plates were incubated for 24 hours at 37°C in 5% CO2. After removing the cells from the upper surface of the membrane with a cotton swab, cells on the lower surface were stained with crystal violet sulfate and counted under a microscope (Olympus, Tokyo, Japan).
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2

Aquatic Ecosystem Monitoring via Multidisciplinary Sampling

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During the summer (June) of 2018 to spring (March) of 2020, 192 water samples in the experimental area and the control area of the artificial structures were collected in four different depths (0, 2, 4 and 6 m) in 8 quarters (Fig. 1). Each of the two groups had 5 randomly repeated sampling points which include AS1-AS5 and CW1-CW5. For each sample, 1.0 L of water was taken from the fixed depth of the water layer using a sterile bottle, and the sterile bottle containing the water sample was immediately placed on the ice [52 ], and then filtered through 0.22 μM the porous polycarbonate membrane (Millipore, MA, USA) by Shimadzu vacuum membrane pump (VP–10L). The samples were stored in liquid nitrogen until DNA was extracted. Temperature, pH, DO, Chlorophyll-a and total dissolved solids were measured by Macro-900 (Palintest, Germany) handheld multi-parameter field instrument (Additional file 1: Table S5). A total of 192 water samples were collected, and about 1000 ml of each water sample was used to measure the physicochemical factors by Macro-900. All fish were sampled using multimesh gillnets that were 18 m long, 1.5 m height with mesh sizes between 6.25 and 60 mm of the following order: 45, 20, 6.25, 10, 55, 40, 12.5, 25, 15, 60, 35, and 30 mm [53 ]. All fish were then anesthetized using the 40 mg/L of tricaine methanesulfonate (MS-222) for subsequent sampling.
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3

Atomic Force Microscopy of Fungal Conidia

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Performed in deionized water using a Fastscan Dimension Icon (Bruker Corporation, Santa Barbara, CA). Images were recorded in peak force tapping mode using oxide sharpened micro-fabricated Si3N4 coated with Au cantilevers with a nominal spring constant of ~0.24 N/m (Bruker Corporation). Conidia were harvested from 12–15 days old malt agar slants maintained at ambient temperature, washed extensively with Tween-water, followed by washing twice with MilliQ water, subjected to PFA-fixation (Aimanianda et al., 2009 (link)) and then immobilized by mechanical trapping into porous polycarbonate membranes (Millipore). After filtering a concentrated conidial suspension, the filter was gently rinsed with deionized water, carefully cut, attached to a steel sample puck using a small piece of double face adhesive tape, and the mounted sample was transferred into the AFM liquid cell while avoiding dewetting. Data processing was performed using the commercial Nanoscope Analysis software (Bruker Corporation). At least eight conidia were analyzed to determine the percentage of conidia completely and partially covered by the surface rodlet-layer.
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4

Bacterial Adhesion Quantified by AFM

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Gold cantilevers (OMCL-TR4, Olympus Ltd., Tokyo, Japan) with a nominal spring constant of ~0.02 N×m-1 were functionalized with fibrinogen using thiols chemistry as previously described for substrates. The spring constants of the cantilevers were measured using the thermal noise method. Bacteria from exponential phase culture were immobilized by mechanical trapping into porous polycarbonate membranes (Millipore, Billerica, USA) with a pore diameter of 0.8 μm. After filtering a cell suspension, the membrane was rinsed with PBS, cut into piece (1 x 1 cm2) and attached to a steel sample puck using a small piece of double-face adhesive tape. The mounted sample was transferred into the AFM liquid cell while avoiding de-wetting. Bare tips were first used to localize and image individual cells and then replaced by functionalized tips. Adhesion maps were obtained by recording 32-by-32 force-distance curves on areas of 500 x 500 nm2, using an applied force of 250 pN, a constant approach-retraction speed of 1 μm×s-1 and a contact time of 100 or 500 ms. Data were analyzed using the Nanoscope software from Bruker (Santa Barbara, USA). Adhesion forces were calculated considering the last peak for each curve and adhesive events are displayed as light pixels. For each condition, experiments were repeated for at least 3 times with independent cultures.
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5

AFM Imaging of Caulobacter Cells

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Caulobacter cells grown overnight in liquid PYE were rinsed in PBS buffer and resuspended in 4% paraformaldehyde (Sigma-Aldrich) solution for 1 hr at room temperature for fixation. Cells were then rinsed in PBS buffer and filtered through polycarbonate porous membrane (Millipore, Billerica, MA, pore size: 3 µm). AFM imaging was performed using a Nanoscope VIII Multimode (Bruker Corporation, Santa Barbara, CA) and oxide-sharpened microfabricated Si3N4 cantilevers with a nominal spring constant of ∼0.01 N/m (Microlevers, Veeco Metrology Group). After filtering the cell culture, the filter was gently rinsed with the buffer, carefully cut (1 cm × 1 cm), attached to a steel sample puck using a small piece of double face adhesive tape, and the mounted sample was transferred into the AFM liquid cell while avoiding dewetting. Images were taken in PBS buffer in contact mode under minimal applied force. Images were analysed using Nanoscope 8.10 software (Bruker, Santa Barbara, CA). Rms (root mean square) roughness values were calculated on 250 × 250 nm2 areas of the high magnification height images subjected to second order filtering.
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6

Atomic Force Microscopy Imaging of Biofilms

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For imaging in air, AFM contact mode images were obtained at room temperature using a Nanoscope VIII Multimode AFM (Bruker Corporation, Santa Barbara, CA) with oxide-sharpened microfabricated Si3N4 cantilevers with a nominal spring constant of 0.01 N m–1 (MSCT, Bruker Corporation). Force data were analyzed using the Nanoscope software (version 8.15, Bruker) and Matlab software (version R2013b). One hundred microliters of biofilm-induced cells was put in contact with freshly cleaved mica supports mounted on steel pucks. The samples were incubated for 2 h at 37 °C, gently rinsed in three successive baths of ultrapure water (Elga, purelab water), and allowed to dry at 30 °C overnight. For imaging in liquid, bacteria were immobilized by mechanical trapping into a polycarbonate porous membrane (Millipore) with a pore size similar to the cell size. After the cell suspension was filtered, the filter was gently rinsed in M63, carefully cut (1 cm × 1 cm), attached to a steel sample puck using a small piece of double-faced adhesive tape, the mounted sample transferred into the AFM liquid cell while avoiding dewetting, and imaged under minimum applied force using MSCT cantilevers.
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