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. Author manuscript; available in PMC: 2024 Dec 2.
Published in final edited form as: Methods Mol Biol. 2024;2801:111–124. doi: 10.1007/978-1-0716-3842-2_9

Evaluation of Connexin Hemichannel Activity In Vivo

Francisca M Acosta 1, Manuel A Riquelme 1, Sumin Gu 1, Jean X Jiang 1
PMCID: PMC11610261  NIHMSID: NIHMS2036347  PMID: 38578417

Abstract

Connexin hemichannels (Cx HCs) are hexameric structures at the cell plasma membrane, whose function as membrane transport proteins allows for the passive flow of small hydrophilic molecules and ions (≤1 kDa) between the cytosol and the extracellular environment. Activation of Cx HCs is highly dependent on pathological conditions. HC activity provokes changes in the microenvironment, inducing the dissemination of signaling molecules in both an autocrine and paracrine manner. Given the elicitation of a variety of signaling pathways, and assortment of Cx species and dispersion throughout the body, Cx HCs have been implicated in a range of processes such as cell proliferation, differentiation, cell death, and tissue modeling and remodeling. While studying the expression and localization of Cx HCs can be done using traditional laboratory techniques, such as immunoblot analysis, measuring the functionality/activity of the HCs requires a more explicit methodology and is essential for determining Cx-mediated physiological changes. The study of Cx HC function/activity has focused mainly on in vitro measurements through electrophysiological characterization or, more commonly, using HC-permeable dye uptake studies. Here, we describe the use of dye uptake to measure Cx HC activity in vivo using mechanically stimulated osteocytic Cx43 HCs with Evans blue dye as our model.

Keywords: Connexin hemichannel, Dye uptake, Pathology, In vivo, Cx43

1. Introduction

Hemichannels (HCs) from two adjoining cells can dock head-to-head, forming a gap junction channel that mediates cytoplasmic communication, while undocked HCs can also serve as sovereign communication conduits for extracellular communication between the cytosol and the extracellular environment [1, 2]. A connexin (Cx) HC is formed by six Cx monomers. In humans, 21 Cx species have been identified distributed in almost all organs and cell types, with some exceptions being red blood cells and healthy mature skeletal muscle myofibers [2]. Members of the Cx family are named based upon their calculated molecular weight; for example, connexin 43 (Cx43), the most abundant connexin isoform, has a molecular weight of 43 kDa [2].

Autocrine and paracrine cellular communication mediated by Cx HCs facilitates the diffusion of ions and metabolites. HCs are permeable to ions, such as sodium (Na+), potassium (K+), calcium (Ca2+), and signaling molecules ≤1 kDa, such as adenosine triphosphate (ATP), prostaglandin E2 (PGE2), glutamate, and other small molecules such as dyes with low molecular weight [3, 4]. Under normal physiological conditions, undocked Cx HCs remain mainly closed; however, pathophysiological conditions, such as during injury or disease, drive their opening, promoting the extracellular release of HC permeable molecules, thus they are known to be major regulators of cellular homeostasis and disease [5]. Among the mechanisms underlying Cx HC activation are alkaline pH, hypoxia, metabolic inhibition, oxidative stress, temperature, and low extracellular Ca2+ [6-8]. Practices such as membrane depolarization, mechanical loading, the introduction of cytokines/growth factors, or deliberate increase of intracellular calcium concentration are also means of stimulating Cx HC opening [9-11].

Given this perspective, when studying Cx HCs in the context of pathophysiological development and cellular homeostasis, beyond the measurement of Cx HC expression and localization, evaluation of HC activity/function is critical. These analytical measurements are commonly done through the use of electrophysiological characterization or, more commonly, using HC-permeable dye uptake studies [3, 12]. In particular, dye uptake has been broadly used as a means of measuring in vitro plasma membrane permeability with charged dyes, such as ethidium bromide, DAPI, Lucifer yellow, Evans blue, among others. These dyes, in conjunction with the addition of specific blockers, could be used to measure the partial participation of Cx HC activity in the increase of membrane permeability [13-15]. The principle behind this technique is that these charged hydrophilic fluorescent dyes can be added in high concentrations extracellularly and cannot innately cross the plasma membrane. In the presence of stimulation, activating or opening HCs, like those mentioned above, the dye diffuses into the cytoplasm through functional and permeable Cx HCs. Excess extracellular dye is then rinsed off, leaving behind dye trapped in the interior of the cells. The incorporated dye can then be quantified using fluorescence microscopy [14]. Explicitly, images containing different fields of view are captured and quantified, utilizing the pixel fluorescence intensity of the dye captured, within the cells of interest, as an index of an increase in the plasma membrane permeability, or Cx HC activity [14].

This same approach, utilizing dye uptake as a means of measuring Cx HC activity/function, can be transferred to measure in vivo Cx HC activity [13, 16-18]. This powerful technique, when coupled with Cx specific inhibitors, mice models with genetic ablations, or different stimulatory techniques, can be used to study not only the activity/function of these HCs using a physiologically relevant approach, but also couple this with local and global changes in pathophysiology. Here, we demonstrate this method using mechanically stimulated Cx43 HCs, in osteocytes, with Evans blue dye as our model. The advantage of using this model is that bone osteocytes primarily express Cx43 and unlike gap junction channels forming at narrow tips of dendrites, Cx43 HCs are richly present on the osteocyte cells surface [19, 20].

Cx43 HC activity was analyzed following (1) intravenous injection of Evans blue dye in mice; (2) activation of HCs by tibial cyclic axial compressions; (3) intracardiac perfusion and tissue fixation; (4)collection of tibial bones, processing, and cryosectioning; and (5)measurement of the fluorescence intensities of osteocytes, indicative of dye infiltration and HC activity, using NIH ImageJ software (Fig. 1).

Fig. 1.

Fig. 1

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Overview of process for in vivo quantification of Cx HC activity, starting at the point of postmodel development

2. Materials

Mice utilized for the study should ideally contain groups with potential differences in Cx HCs including the treatment with Cx HC specific inhibitors, mouse models containing genetic ablations, and/or the utilization of different stimulatory techniques, in order to obtain quantifiable Cx HC activity changes. For the example data presented here, Cx43 HCs in osteocytes were the target and mechanical stimulation, through tibia loading, was utilized in WT and transgenic mice (R76W and Δ130–136), driven by a 10-kb DMP1 promoter with the overexpression of dominant negative Cx43 mutants in osteocytes. The R76W is a dominant negative mutant that blocks the endogenous gap junction channel, but not the HC function, and the dominant negative Δ130–136 mutant inhibits the activity of both types of channels [21]. Using these models, we have reported differences in bone anabolic function in response to mechanical loading/Cx43 HC activity, which we quantified utilizing this methodology [18].

2.1. Evans Blue Dye Preparation, Injection, and Collection

  1. Dissolve 20 mg/mL Evans blue (EB) dye in sterile saline solution. Solution should be filtered with a 0.22 μm filter to remove any particulates (see Note 1).

  2. 0.22 μm filter.

  3. Scale.

  4. Antiseptic solution (e.g., 70% ethanol (EtOH)).

  5. Gauze pads.

  6. 0.5 mL 25G insulin syringe.

  7. Warmed phosphate-buffered saline (PBS) pH 7.4.

  8. Mice mechanical restraint device.

  9. Isoflurane.

  10. Precision vaporizer system with, oxygen, anesthesia mask, and induction chamber.

  11. 4% paraformaldehyde (PFA).

  12. Surgical scissors, forceps, and hemostats.

  13. Blunt 15G perfusion needle.

  14. 3 × −100 mL syringe.

  15. Peristaltic pump (optional).

2.2. Tibial Mid-diaphyseal Strain Measurements and Cyclic Tibial Loading

  1. Strain gauge.

  2. Bose ElectroForce LM1 test instrument.

  3. Bridge completion module.

  4. Signal conditioner/amplifier systems.

  5. Tibial loading Device (Custom Apparatus).

  6. Isoflurane.

  7. Precision vaporizer system with oxygen, anesthesia mask, and induction chamber.

2.3. Cryosectioning, Imaging, and Quantification

  1. 10% ethylenediaminetetraacetic acid (EDTA) in PBS.

  2. 15% sucrose in PBS.

  3. 30% sucrose in PBS.

  4. Optimum cutting temperature compound (OCT).

  5. Liquid nitrogen.

  6. Cryostat.

  7. Hydrophobic Barrier PAP Pen.

  8. 4′,6-diami-dino-2-phenylindole (1 μg/mL, DAPI).

  9. Fluoromount-G.

  10. Optical fluorescence microscope.

  11. NIH ImageJ software.

3. Methods

Carry out all procedures at room temperature unless otherwise specified. All animal protocols should be performed following the National Institutes of Health guidelines for care and use of laboratory animals and approved by Institutional Animal Care and Use Committees (IACUC).

3.1. Tibial Mid-diaphyseal Strain Measurements

  1. (see Note 2) Complete this step prior to experiment set-up to ensure that proper calculation is made prior to study, to determine the applied load (N) required to generate equivalent micro strains (με) between your testing groups.

  2. Euthanize mice, from each testing group, with isoflurane.

  3. Exposed tibia bones using surgical instruments (see Note 3).

  4. Attach strain gauge to the tibial diaphyseal anterior-medial surface (~37% down the tibia bone from the proximal end).

  5. Using an electromagnetic testing system, such as the Bose ElectroForce LM1 with an attached bridge completion model and a conditioner/amplifier system, apply dynamic compressive loads from 0 to 9.5 N at the ends of the tibia.

  6. Plot the slope between the load and micro strain (N/με), (R2 > 0.99), indicative of the “Tibial Compliance” (Fig. 2).

  7. Based on the calculated load–strain relationship, determine the force/load (N) to use for mechanical loading of groups (see Note 4).

Fig. 2.

Fig. 2

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Measurement of tibial mid-diaphyseal strain. (a) Placement of strain gauge in exposed tibial bone. (b) Setup for measurement of tibial compliance with an electromagnetic mechanical testing system

3.2. Evans Blue Dye Tail Vein Injection

  1. Preweigh mice using scale.

  2. According to mouse weight, prepare syringes of EB solution (20 mg/mL in sterile saline) using insulin syringes, dye injected should be 200 mg/kg.
    1. For example, for 20 g mouse, 200 μL of EB solution should be used.

  3. Place mice in mechanical restraint of choice with tail vein protruding.

  4. Wipe tail with antiseptic solution.

  5. Immobilize tail vein with nondominant hand and rotate ~90° to access the lateral tail vein (see Notes 5 and 6).

  6. Hold the tail taut with nondominant hand, near the tip of the tail, and insert the needle into the vein at a minimal angle (≤30°), to tail, with the beveled edge of needle facing upwards (see Note 7).

  7. Advance the needle inward until ~ ½ the length is inside (~5 mm), careful as to not to perforate the vein (see Note 8).

  8. Upon completion, remove needle and gently apply pressure at site of injection until bleeding has stopped (see Note 9).

  9. Place animal back into housing container for 20 min then immediately proceed with tibial cyclic axial loading.

  10. Repeat with any additional mice, as needed (Fig. 3).

Fig. 3.

Fig. 3

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Injection of mice tail vein with Evans blue dye for measurement of permeable Cx HCs. (a) Setup with mice in mechanical restraint with tail protruding and accessible for injection. (b) Anatomical landmarks needed for localization of tail vein

3.3. Tibial Cyclic Axial Loading

  1. Ensure proper weights are attached to the loading device based on calculated load–strain relationship (see Note 10), in addition to attaching a 0.4 N continuous static preload to hold tibia in place.

  2. Preanesthetize mice by using a precision vaporizer system connected to an oxygen tank and placing mice inside the induction chamber and seal/close top for ~2–5 min at 2.5% isoflurane in 100% oxygen.

  3. Ensure that animal has reached a surgical plane of anesthesia via toe pinch.

  4. Switch system flow and transfer mice to an anesthesia mask/nose cone in proximity to loading apparatus and adjust gas flow with vaporizer to 2–3% isoflurane.

  5. Position the tibia to be loaded (see Note 3) into loading apparatus with an upper padded cup containing the knee connected to loading device, conducting the compressions, and a lower cup holding the heel in place.

  6. Conduct the cyclic axial compressive loading (1200 cycles (10 min) at 2-Hz frequency with a sinusoidal waveform).

  7. Place animal back into housing container for 40 min then immediately proceed with euthanization and perfusions.

  8. Repeat with any additional mice, as needed (Fig. 4).

Fig. 4.

Fig. 4

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Mice in vivo axial loading for activation of osteocyte Cx43 HCs. (a) Diagram of tibia positioning in the loading device and blue arrow demonstrating the direction of loading. (b) Schematic of the custom loading setup for tibial cyclic axial loading, specifically showing anesthetized mice undergoing loading in the left tibia using a predetermined force, to equalize strain

3.4. Euthanization and Perfusion

  1. Euthanize mice with isoflurane.

  2. Place the mouse on its back and firmly tape down its four paws to immobilize.

  3. Using the surgical tools to perform a thoracotomy, cutting above the diaphragm and along the ribs up to the collarbone, to expose the heart, taking special care not to puncture the heart.

  4. Carefully cut away any other tissues that may be occluding the heart to provide a clear view of the heart and vessels.

  5. Connect a 15G blunt perfusion needle to a 100 mL syringe filled with PBS and flush the tubing and needle to expel air (see Note 11).

  6. Pass the blunt 15G perfusion needle filled with PBS, through the tip of the left ventricle (see Note 12) at an angle parallel to the midline of the heart until it enters the ascending aorta, visibly seen through aorta wall, but not reaching the aortic arch.

  7. Using a hemostat, clamp the heart, to secure the needle and prevent leakage.

  8. Make an incision on the right atrium using a small scissor to allow for the flow out of blood/liquid being perfused.

  9. Begin PBS perfusion, to remove extraneous EB, by injecting warmed PBS at a constant speed of 4–5 mL per min for 10 min by manually pushing the syringe or using a peristaltic pump.

  10. Switch from PBS to 4% PFA, making sure no bubbles get into the perfusion system, and perfuse mice, as before, to fix tissues.

  11. For the last perfusion, switch from 4% PFA back to PBS, to completely administer the remaining PFA in the system, and perfuse as before (see Note 13).

  12. Immediately proceed with isolation and processing of tibias.

  13. Repeat with any additional mice, as needed (Fig. 5).

Fig. 5.

Fig. 5

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Transcardiac perfusion of mice for tibial bone fixation. (a) Anatomical landmarks needed to perform procedure and position and direction of needle insertion. (b) Demonstration of the clamping of the needle to prevent movement and leakage. (c) Small incision being made in the right atrium to allow for the outflow of perfused liquids. (d) Flowchart of perfusion steps

3.5. Isolation, Processing of Tibias, and Cryosectioning

  1. Using the surgical instruments, isolate tibias taking special care in removing surrounding muscle and connective tissue.

  2. Fix the tibias with 4% PFA for 2 days in 4 °C (in 10× tissue volume).

  3. Rinse tibias with PBS three times for 30 min each.

  4. Decalcify tibias with 10% ethylenediaminetetraacetic acid (EDTA) for 3 weeks (in 10× tissue volume) at 4 °C, changing out EDTA two times a week.

  5. Following decalcification, rinse tibias with PBS three times for 5 min each.

  6. For cryopreservation, immerse decalcified tibias in 15% sucrose for 12 h at room temperature then 30% sucrose overnight at 4 °C (in 10× tissue volume).

  7. Embed tibias longitudinally in OCT compound.
    1. Dip the embedding mold repeatedly in liquid nitrogen for a short time until the OCT changes to a porcelain white color, indicating freezing of sample. Store at −20 °C for at least 12 h before cutting sections. Long-term storage can be done at −80 °C for ≤1 month (see Note 14).

  8. Using a cryostat, set to −20 °C, cut longitudinal frozen tissue sections of 12 μm thickness, and place in a − 20 °C freezer until ready for staining.

3.6. Tibia Staining and Imaging

  1. Remove slides from freezer and let samples come to room temperature for ~30 min.

  2. Immerse the frozen tissue sections in cold acetone, which increases the permeability and fixation, at −20 °C for 2 min, and then air-dry the sections.

  3. Use Hydrophobic Barrier PAP Pen to draw a liquid barrier around each tissue section.

  4. After drying, rinse the slide with PBS, to remove residual OCT, washing three times, 5 min for each time.

  5. Counterstain sample with DAPI for 10 min, followed by a PBS wash three times, 5 min each time.

  6. Mount slides with Fluoromount-G and seal with a cover slide (keep tissue hydrated throughout this process).

  7. Store the slides in a black plastic box at 4 °C to prevent quenching of fluorescence signals until imaging is completed.

3.7. Osteocyte Cx43 HC Activity Imaging and Quantification

  1. Begin imaging, using an optical fluorescence microscope, by first standardizing image capture, starting with the slides with the cells expected to produce the brightest signals (the group with the expected highest Cx HC activity) and fix exposure time for comparisons between groups (see Note 1).

  2. Capture images by identifying osteocytes in the cortical bone area located at ~37% of the tibial diaphysis.

  3. Using the NIH ImageJ software (Fiji if preferred) begin the analysis by preparing images for analysis by first succumbing images to batch conversion into appropriate file format and enhancement, if needed (see Note 15).

  4. Using this same ImageJ software, quantify the fluorescence intensity of EB within osteocytes making sure to do a minimum of 20 cells per section (three sections per mouse) (see Note 16).

  5. Once all images have been analyzed, an ANOVA/t-test can be used to determine statistical significance (Fig. 6).

Fig. 6.

Fig. 6

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Imaging and quantification of in vivo osteocyte Cx43 activity. (a) Imaging of mice with osteocyte Cx43 HCs activated ((+) Loading) and nonactivated ((−) Loading) in the cortical bone in the tibial diaphysis. Callouts, highlighting the ability to visualize EB uptake (red) in the cytoplasm of osteocytes with activated/functional HCs. Only WT mice shown. (b) Quantification of EB uptake, demonstrating the level of permeable osteocyte Cx43 HCs, synonymous to HC activity. Here, WT and mutant R76W mice have functional osteocytic Cx43 HCs, while the mutant Δ130–136 mice lack active HCs, thus see no significant increase of EB uptake [21]

4. Notes

  1. Cx HCs are permeable to Evans blue (EB) (Mw 960.81 Da), which possesses four negative charges and it is a fluorescent dye with two excitation wavelengths λx) at 470 nm and 540 nm and a peak emission (λem) at 680 nm, producing fluorescence in the red to far-red spectrum. More importantly, this molecule can be fixed by fixative reagent to allow the detection on the tissue sections [22]. Because of low cell permeability to EB, it has been used to evaluate cell viability. EB can bind to albumin protein, so it has also been used to tract the integrity of blood barriers such as the blood–brain barrier [23]. It has also been reported that EB blocks the activity of glutamate receptors and the transport and activity of P2 receptors [24-26]. Pannexin 1 are high-capacity channels that share similar permeability and pharmacology with Cx HCs. However, it has been shown that they are impermeable to EB [13]. Of note, other small fluorescence dyes can be used, however ensure the dyes can be fixed on tissue sections, prior to adoption for this application.

  2. This protocol of the relationship between applied compressive loading and bone tissue deformation of the left tibia was established for 15-week-old mice in vivo based on previously reported protocols [27, 28].

  3. For our experiments, we typically only calculate these metrics/load mice unilaterally, meaning for us the left tibia is measured/loaded and the right tibia is used as a contralateral, nonloaded control.

  4. We typically select peak forces to generate peak periosteal strains of 1200 με at the cortical midshaft since this strain level has been shown to elicit an anabolic response at this region previously [29].

  5. Since warming the mouse/tail causes vasodilation, this technique can be utilized for better vein visibility. This can be done using a heating chamber, heating pad, a heating lamp, or dipping the tail in warm water, prior to injection. However, caution should be taken to avoid overheating animal.

  6. The mouse has two lateral tail veins and a tail artery located on the ventrum of the tail, either the left or right lateral tail veins may be used.

  7. During this step, it is the best practice to do injections close to the tip of the tail since (1) in case first injection is unsuccessful and vein is perforated, you can retry by advancing upward, and (2) the vein is deeper the closer you move toward the body and can be more difficult to access.

  8. If injection is successful, blood should flash to the hub of the needle, vein will lighten, and liquid will flow easily during administration. No resistance should be felt. Visibly, mice will also begin to change to a blueish color, the snout, paws, and tail will turn blue, indicating successful administration within ~5 min.

  9. A clotting agent, such as styptic powder, silver nitrate sticks, or cautery pen, can be used to assist with the stoppage of blood flow.

  10. The cyclic axial compressive load being applied to the left tibia of each mouse is done using a custom loading device based on previous studies [27, 28]. Keep in mind 1 gram (g) = 0.0098 Newton earth (N).

  11. Blunt needed is optional, but preferred, to reduce the risk of puncturing the aorta. Blunting of the needle can be done manually using an assortment of tools/sandpaper. Additionally, needle of bigger gauges/not perfusions based, at the preference of the investigator.

  12. If blunt needle is used and there is resistance when trying to insert needle into the left ventricle, prior to insertion, a very small incision can be made to simplify needle entrance. It is recommended before injection, the midline and apex of the heart be identified to ensure accurate insertion of needle into left ventricle.

  13. For more detail protocol on perfusion of rodents please see [30, 31].

  14. Take special care to not immerse samples too quickly to avoid “cracking” of tissue samples.

  15. Though image conversion/enhancement can be done (example see: [32]) ensure images are not manipulated ununiformly skewing results.

  16. For a more in detail example of image processing/quantification please see [32].

Acknowledgments

This work was supported by NIH CA148724 & NIH F32DK134051 (to F.M.A.), AR072020 (to J.X.J.) and Welch Foundation grant: AQ-1507 (to J.X.J.)

Figures partially created with BioRender.com.

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