Theranostics 2015; 5(1):97-109. Doi:10.7150/thno.10298 Research Paper Theranostic mRNA-loaded Microbubbles in the Lymphatics of Dogs: Implications for Drug Delivery Heleen Dewitte 1*, Katrien Vanderperren 2*, Hendrik Haers 2, Emmelie Stock 2, Luc Duchateau 3, Myriam Hesta 4, Jimmy H.

Saunders 2, Stefaan C. De Smedt 1, Ine Lentacker 1 1. Laboratory for General Biochemistry and Physical Pharmacy, Faculty of Pharmacy, Ghent University, Ghent, Belgium; 2.

Jan 20, 2008. 4.1.5 Scalpel. No specific vendor required. CATEGORY: Laboratory Work. SOP Number: Lymph Node_Rutgers. TITLE: Separation and. Coulter Allegra 6R. Rotor Gh 3.8A or comparable. No specific vendor required. 4.1.9 ViCell XR. Beckman Coulter. 4.1.10 Liquid nitrogen Tank.

Department of Veterinary Medical Imaging and Small Animal Orthopaedics, Faculty of Veterinary Medicine, Ghent University, Merelbeke, Belgium; 3. Department of Comparative Physiology and Biometrics, Faculty of Veterinary Medicine, Ghent University, Merelbeke, Belgium; 4. Department of Animal Nutrition, Faculty of Veterinary Medicine, Ghent University, Merelbeke, Belgium.

* both first authors contributed equally to this work. Table 1 Design of the crossover CEUS study. Day Dogs Microbubble injections Week 1, day 1 1, 2, 3 Left: unloaded microbubbles Right: mRNA-loaded microbubbles Week 1, day 2 4, 5, 6 Left: mRNA-loaded microbubbles Right: unloaded microbubbles 2 week wash-out Week 3, day 1 1, 2, 3 Left: mRNA-loaded microbubbles Right: unloaded microbubbles Week 3, day 2 4, 5, 6 Left: unloaded microbubbles Right: mRNA-loaded microbubbles Image analysis Image analysis was performed using QLAB quantification software (Philips) and ImageJ.

Microbubble migration distances were calculated as a linear distance between the injection site and the most distant microbubble contrast signal. Measurements were performed by two independent, blinded observers. Maximal echo intensities were measured using ImageJ at different timepoints after injection on a region of interest (ROI) within the lymphatics (i.e.

Within the enhanced lymph nodes or within the lymph vessels when no lymph node enhancement could be observed). Analysis of burst destruction of intranodal microbubbles was performed by drawing a ROI over the lymph node of interest, after which the mean echo intensity was studied as a function of time using the QLAB quantification software. In the figures, time is indicated in min:s on the CEUS images, and the depth scale bar is shown at the right-hand side of the corresponding B-mode images (the distance between two ticks is 5 mm in all images). Statistical analysis The migration distance and the maximal echo intensity was compared between the loaded and unloaded bubbles by a mixed model with dog as random effect and period and bubble type as categorical fixed effects, using the F-test at the 5% significance level.

The contrast echo intensity over time was compared between the loaded and unloaded bubbles by a mixed model with dog as random effect and period, time, bubble type and the interaction between time and bubble type as categorical fixed effects, using the F-test at the 5% significance level. Within and between observer variability was determined by estimating the within and between observer variances for a same assessment (i.e., the same dog, side, bubble type and period) by the restricted maximum likelihood procedure (REML). These estimated variances were next used to determine the range in which 95% of the differences between two measurements of the same observers and two different observers are contained.

SAS version 9.3 was used for all analyses. Results Characterization of unloaded and mRNA-loaded microbubbles mRNA-lipoplex loaded microbubbles were prepared by first complexing the mRNA to cationic liposomes, to form mRNA-lipoplexes in which the mRNA is protected against degradation (: Figure S1).

These mRNA-lipoplexes are then attached to the surface of avidinylated lipid microbubbles, as shown in Figure A. To demonstrate effective loading, we performed confocal microscopy on microbubbles that were loaded with fluorescently labeled mRNA-lipoplexes. As demonstrated in Figure B, the mRNA-lipoplexes are clearly present around the microbubble surface, and we did not observe any aggregation. The size and concentration of the microbubble preparations was determined via coulter counter measurements.

The unloaded microbubbles had a mean number diameter of 2.48 ± 1.57 µm. Attachment of mRNA-lipoplexes did not significantly alter the mean diameter (1.93 ± 1.25 µm) ( Figure C).

The concentration of the microbubble preparations was 2.68 ± 0.86 x 10 9 bubbles ml -1. Factors influencing intralymphatic CEUS Based on previous experiments studying the lymphatic drainage of SonoVue TM microbubbles by Goldberg et al. [], we performed subcutaneous injections of both unloaded and mRNA-loaded microbubbles in dogs. When performing these experiments, we observed that a number of factors influenced the CEUS signals. First of all, we noticed important differences in the migration of the microbubbles depending on the location of the injection site. When injecting the microbubbles in the groin, close to the inguinal lymph node, we could not observe any lymphatic uptake of the contrast agents. The reasons for this are probably dual.

First of all, lymphatic uptake occurs through a passive process that depends on the interstitial pressure at the injection site. Hence, regions such as the groin, where the skin is loose and the interstitial pressure is low, are not preferable []. In addition to pressure-related differences, some tissues, such as the mammary regions of female dogs, contain a more developed lymph vessel network. In accordance to this, initial s.c. Injections of the contrast agents in the loose skin of the abdominal region did not result in migration of the contrast agents from the injection site. This despite the fact that the microbubbles were injected at a distance of merely 20 mm from the inguinal lymph node, as indicated by palpation and B-mode ultrasound imaging.

On the other hand, injection around the dog's nipple resulted in extensive microbubble drainage in all female dogs, as shown in Figure. Migration from the injection site was seen for all injections with both unloaded (8/8) and mRNA-loaded (8/8) microbubbles in the female dogs, and after most of the injections around the nipples of the male dogs (4/4 for the unloaded bubbles and 2/4 for the mRNA-loaded bubbles).

This is not unexpected, as the mammary regions accommodate a vast network of lymph vessels that lead to multiple clusters of lymph nodes close to the mammary glands, as well as more distant gastric and mesenteric lymph nodes, depending on the location of the mammary gland (e.g. Axillary versus inguinal mammary glands) []. Based on these observations, further injections were performed s.c.

Around the left and right nipple of the inguinal mammary gland. Figure 2 Influence of animal properties on the lymphatic CEUS images. Pictures show the CEUS images obtained after s.c.

Injection of unloaded microbubbles around the mammary glands of a male dog (A1), a spayed female dog (B1) and an intact female dog (C1). Respective B-mode images are shown in A2, B2 and C2. The injection site is indicated as “i.s.” (the injection site for images C1 and C2 is out of the field-of-view), arrows indicate the accumulation of microbubbles in the lymph nodes and lymph vessels are pointed out with asterisks. Time after microbubble injection is noted on the CEUS images (in min:s). (Click on the image to enlarge.) Besides the effects of the injection site, inter-animal variation in the mammary gland lymphatic network also played a role. In male dogs, the mammary glands are markedly less developed compared to females, which also results in a less dense lymphatic network and fewer intramammary nodes.

Moreover, the male dogs clearly showed a less thick subcutaneous fat layer (i.e. Where the lymph vessels and nodes are located) compared to all females, as evidenced by the B-mode images in Figure, images A2, B2 and C2. As a result, contrast agent migration after injection was less pronounced in the males compared to the females. In the intact male, injection next to the testicles resulted in more distant microbubble migration when compared to injection around the mammary glands, which therefore represented a valuable alternative to injection around the mammary glands. Beside these gender-related factors, we also observed differences between spayed and intact females. After microbubble injection in the intact female, the lymphatic structures were visible as a vast network with multiple nodes and vessels extending more deeply into the subcutaneous fat pad (Figure, images C1 and C2, and: supplementary video 1, observed for all injections with both unloaded and mRNA-loaded microbubbles).

Such a branched lymph network could not be observed in any of the 3 spayed females. Of course, this inter-animal variation makes it more difficult to fully exclude anatomical bias when comparing the lymphatic drainage of unloaded and mRNA-loaded microbubbles. The cross-over design used for this study, where all dogs receive s.c. Injections with both unloaded and mRNA-loaded microbubbles in the nipples of both the left and the right caudal mammary gland, was used to take this variability into account and allow a better comparison of both microbubble types. Lymphatic network visualization After microbubble injection and massaging of the injection site, the migration of the contrast agent was observed unidirectional, with the contrast agents leaving the injection site at only one end in the spayed females and the males, as exemplified by Figure A. In the intact female, microbubble migration occurred both cranially as well as caudally. The migrating contrast agents were followed through the lymph vessels towards draining lymph nodes (Figure, Images B1, B2, C1 and C2, and: supplementary video 2).

Importantly, in none of the injections (0/24) did we observe blood pool contrast enhancement, indicating that the microbubbles were restricted to the interstitium, the lymph vessels and the lymph nodes. Moreover, when the contrast agents were injected on one side (either left or right) of the animal, contrast signal was only observed on that same side of the animal.

We did not observe contrast crossing over the midline of the dogs in any of the cases. One or multiple lymph nodes could be identified using CEUS after 9/12 injections of both unloaded and mRNA-loaded microbubbles, and the size of the enhanced lymph nodes ranged from 1.6 to 7.0 mm.

Interestingly, CEUS imaging revealed rather detailed information on the lymphatics' anatomy: in accordance to previous reports, CEUS could provide information on the location of the lymph nodes, the number of afferent and efferent lymph vessels []. In addition, we could observe different patterns of contrast agent presence within the nodes. Some lymph nodes, such as the one shown in Figure A, are completely filled with contrast agent, whereas others rather exhibit a “hollow” appearance, with contrast material only appearing at the outer rim of the node, as shown in Figure, images B1, B2, C1 and C2. This can be explained by the fact that afferent lymph vessels can either directly discharge their content into the draining lymph nodes, or the afferent lymph vessels run through or over the nodes, without effectively discharging the lymph within the node []. Figure 3 Contrast agent migration from the injection site into the lymph vessels and nodes.

Upon microbubble injection, unidirectional transport of microbubbles away from the injection site, through an afferent lymph vessel into a draining lymph node can be observed with CEUS (A1). Image (B1) shows the trafficking of microbubble contrast signal through branched lymph vessels. In (C1), the CEUS image shows a lymph node connected to one afferent lymph vessel and 3 efferent lymph vessels that take the contrast agents further away from the injection site. Corresponding B-mode images are shown in (A2), (B2) and (C2), respectively. Images were obtained after injection of mRNA-loaded microbubbles in different female dogs.

The injection site is marked as “i.s.” (unless outside of the field-of-view), arrows indicate lymph nodes and asterisks point out lymph vessels. Time after microbubble injection is noted on the CEUS images (in min:s). (Click on the image to enlarge.). Figure 4 Lymph node anatomy observed by CEUS. CEUS can be used to identify the relation between afferent lymph vessels and draining lymph nodes.

Lymph vessels either distribute their content within the lymph node (“filled” nodes, A1-A2) or they go around the lymph nodes without discharging its contents into the node (“hollow” nodes, B1-B2 and C1-C2). In the latter scenario, we always observed a node with a “hollow” appearance (indicated as “H”), followed by a more distant “filled” node (indicated as “F”).

Images were obtained from 3 different animals. Where possible, the injection site is pointed out as “i.s.”. Time after microbubble injection is noted on the CEUS images (in min:s). (Click on the image to enlarge.) Lymphatic CEUS using mRNA-loaded microbubbles versus unloaded microbubbles In this study, we compared the CEUS images after s.c.

Injection of unloaded and mRNA-loaded microbubbles, using three main scoring criteria: (a) the migration distance of the contrast agents from the injection site within the lymphatics, (b) the maximal intensity of the contrast signal and (c) the stability of the contrast agents in the lymph vessels and nodes over time. With regards to the first parameter, the migration distance, we could detect microbubble migration from the injection site for all injections with unloaded microbubbles (12/12), and in 10/12 cases after mRNA-loaded microbubble injections. The 2 injections that did not result in contrast agent drainage were observed in the male dogs (1 in the castrated male and 1 in the intact male). The microbubble migration distance (calculated linearly from the injection site) was on average 24.7 ± 13.7 mm for unloaded microbubbles and 29.0 ± 19.0 mm for mRNA-loaded microbubbles ( Figure A). No significant difference was observed for migration distance between the loaded and unloaded bubbles, with a mean difference equal to 4.33 (95% CI: [-4.55; 13.22]). It should be noted that even though these results were analyzed based on injections of unloaded and mRNA-loaded microbubbles within the same mammary gland, variation based on the exact injection site cannot be excluded, even though injections were always performed caudally from the nipple by the same person. For instance, it was shown by Goldberg et al.

[] that injection sites that were merely 1 cm apart in distance, could result in drainage to a different lymph node, which can obviously impact the microbubble migration distance that was observed. As the exact injection site was not marked at the time of the first imaging session, variation related to the injection site location cannot be fully excluded. The variation between repeated observations of the same observer was 23.9, and the extra variation due to different observers was 9.0 for loaded bubbles, leading to somewhat larger 95% intervals for differences between two measurements of two different observers as compared to the same observer. On the other hand, for the unloaded microbubbles, the variation due to repeated observations of the same observer was 16.7, and there was no extra variation due to different observers, resulting in the same 95% intervals for differences between two measurements of two different observers as compared to the same observer (: Figure S2). In addition to distance measurements, we compared the maximal echo intensity that we could obtain within the lymphatics after injection of unloaded and mRNA-loaded microbubbles. As for the migration distance, we could observe no significant differences in lymphatic contrast enhancement between both types of bubbles ( Figure B), with a mean difference between loaded and unloaded bubbles equal to -2.56 (95% CI:[-15.04; 9.91]). Moreover, when the contrast echo intensity was evaluated as a function of time, we could observe relatively long-term contrast agent stability within the lymphatics, as in all cases where contrast migration was observed, the microbubbles could still be clearly detected 6min after microbubble injection (: Supplementary video 3).

There was merely a slight decay of the echo intensity over this period, and we could not detect significant differences between both types of microbubbles ( Figure ). Keeping in mind the final aim of the mRNA loaded microbubbles, namely the ultrasound-triggered delivery of mRNA to DCs that reside within the lymph nodes, we tested whether intranodal mRNA-loaded microbubbles could indeed be imploded by applying higher-intensity ultrasound bursts.

For this, we used the scanner's preset burst function, and looked at the decrease in mean echo contrast intensity within the lymph node. As shown in Figure, with each burst, the microbubble echo intensity was reduced, reaching a minimum after 6 bursts. A video of this burst destruction can be found in supplementary data (: supplementary video 4).

Discussion In recent years, drug- and gene loaded microbubbles emerged as interesting theranostics after intravenous injection. This, however, is the first report on the development of theranostic mRNA-loaded microbubbles for lymphatic imaging and lymph node detection after s.c. When taken together with our previous research on transfections with these microbubbles, this paves the way for ultrasound-guided microbubble-assisted drug delivery to intranodal cells. To explore this, we aimed to compare unloaded and mRNA-loaded microbubbles with respects to the lymphatic uptake and contrast enhancement in the lymph nodes upon s.c.

Injection in dogs. Figure 7 Burst destruction of intranodal microbubbles. After visualization of unloaded microbubble drainage, the scanner's burst function was used to destroy the microbubbles within the lymphatics. Images show (A) CEUS image with a ROI drawn around a lymph node filled with mRNA-loaded microbubbles and (B) the corresponding B-mode image. Burst analysis is represented in (C) as the echo mean (dB), where each burst is visible as an echo mean peak, followed by a reduction of the contrast echo mean. Time after mRNA-loaded microbubble injection is noted on the CEUS images (in min:s).

(Click on the image to enlarge.) The lipid microbubbles used in this study could be efficiently loaded with mRNA-lipoplexes, and had mean diameters around 2.5 µm, which is comparable to the size of commercially available microbubbles. Lipoplex-loading did not significantly alter the mean microbubble diameter.

Our imaging results indicated that despite animal-related variation, both types of microbubbles could be detected within the lymph nodes within 1min after s.c. Injection around the mammary glands. How exactly the microbubbles are transported from the interstitium to the lymph vessels remains controversial, as it goes against the current view of ideal particle properties for lymphatic uptake. It is generally considered that particles between 10 and 100 nm in size result in the best lymphatic uptake []. Although the appearance of 1 µm particles in the draining lymph nodes was also observed, their uptake efficiency and speed was significantly reduced and a large fraction of the injected particles were retained at the injection site []. In the case of our microbubbles we could observe fast and extensive microbubble uptake into the lymph vessels and draining nodes, despite their larger size. The rapid uptake of microbubbles into the lymphatic system is not well understood.

A first explanation could be found within the other physicochemical properties of the microbubble contrast agents we injected. Besides particle size, lipophilicity and the presence of surface modifications that give stealth properties to injected particles, were also reported to positively impact their lymphatic drainage []. Therefore, the presence of the lipid coat in both the microbubbles and the mRNA-lipoplexes could aid to promote microbubble uptake into the lymph vessels. In addition, the microbubbles as well as the mRNA lipoplexes contain lipids that are conjugated to polyethylene glycol (PEG, 15 mol.% and 2.5 mol.% for bubbles and lipoplexes respectively), which is known to protect the particles against uptake by phagocytes, thus reducing their premature clearance, resulting in enhanced drainage to the lymphatics. On the other hand, Goldberg et al. Proposed a cell-mediated uptake mechanism after s.c. Injection of Sonazoid TM in pigs [].

They performed electron microscopy on isolated lymph nodes after contrast agent injection, and observed the presence of vacuoles in the intranodal phagocytic cells, which could imply that the microbubbles are first ingested by cells, which then transport them from the interstitium to the lymph vessels and draining lymph nodes. However, this cellular transport through the lymphatics is only likely for microbubbles that are easily phagocytosed.

A study by Yanagisawa pointed out that microbubble phagocytosis by primary liver cells was highly dependent on the microbubble composition. Indeed, the authors demonstrated that over 99% of the Sonazoid microbubbles were rapidly phagocytosed by primary liver cells, which can likely be attributed to their shell composition []. Sonazoid TM consists of egg phosphatidyl serine, which is recognized by macrophages and thus enhances phagocytic uptake [, ]. In contrasts, anionic Imavist TM microbubbles (where the anionic charges reduce microbubble contact with cell membranes) and PEGylated SonoVue TM microbubbles were merely for 0% or 7.3% phagocytosed, respectively. As our microbubbles are also PEGylated (by inclusion of 15% DSPE-PEG (3400)-biotin), and no specific targets for macrophage recognition are present, it is unlikely that our microbubbles were carried into the lymphatics inside of cells. Injection, we could observe clear migration of both unloaded and mRNA-loaded microbubbles from the injection site into the lymph vessels and the draining lymph nodes.

Moreover, our CEUS images nicely correspond to previous reports on the lymphatic drainage of microbubbles, which indicates that the observed structures were indeed lymph nodes [-,, ]. The imaging options that are associated with these mRNA-loaded microbubbles could have implications for mRNA delivery as well as for diagnostic purposes. Firstly, the mRNA-loaded microbubbles could provide information on the anatomical features of draining lymph nodes. To our knowledge, this is the first report where CEUS was shown to discriminate between the two different types of connection of lymph vessels to lymph nodes (i.e. The “hollow” nodes and the “filled” lymph nodes) [].

The impact of this might be dual. First of all, from our perspective of future mRNA delivery to intranodal DCs, it would be a major advantage to be able to discriminate between situations where the afferent lymph vessels discharge the mRNA-loaded bubbles within the core of the lymph node (“filled” nodes) versus lymph vessels that merely run over the surface of the lymph nodes, thus bypassing the actual node, resulting in the absence of a microbubble signal in the center of the node (“hollow” nodes). Only in the first scenario will the mRNA-loaded bubbles be able to reach the intranodal DCs, and is DC transfection with mRNA via sonoporation possible. Therefore, delivery of high-intensity ultrasound pulses to induce microbubble implosion and mRNA delivery should only be performed in the lymph nodes that exhibit the “filled” appearance. Other than for ultrasound-guided drug delivery, this anatomical information could also have benefits with regards to SLN detection.

In a tumor setting, the SLN will be the first lymph node that encounters material that directly originates from the tumor, such as disseminating tumor cells. However, if the first lymph node after the tumor has the “hollow” characteristics, this means that the tumor cells do not enter within the node, making it less likely that this node would be populated with tumor cells []. Therefore, lymph node biopsies to detect lymph node metastasis are best performed in the first node in which the content of the tumor-draining lymph vessels are actually discharged. CEUS-guided identification of the first “filled” SLN could therefore have an added value to reduce chance of false negatives in SLN biopsies. Of note, other imaging possibilities such as 3D CEUS imaging, could enable improved visualization of the lymphatic drainage and improve variability in measurements.

When comparing mRNA-loaded microbubbles with unloaded microbubbles, we could not observe statistically significant differences with regards to migration distance, mean contrast intensity and the stability of both contrast agents in the lymphatics. As the size distribution of both types of bubbles did not significantly differ, this was not entirely unexpected. In addition, experiments on similar nanoparticle-loaded lipid microbubbles by Luan et al. Demonstrated that the shell elasticity of individual unloaded and liposome-loaded microbubbles was nearly the same []. The effects of liposome-loading that was observed by the authors were mainly on the microbubble shell viscosity (which was higher for liposome-loaded microbubbles) and thus on the ultrasound-induced microbubble vibrations. Taken together, the impact on these acoustical differences between two types of bubble populations is expected to be limited with respect to contrast-enhancement.

However, it is likely that for molecular imaging and therapeutic purposes, which often involve only a single microbubble or a few microbubbles in a given volume, these differences will need to be addressed: liposome-loaded microbubbles exhibited a higher pressure threshold for microbubble vibration, which might indicate that the pressures that will be needed to implode these bubbles in order to locally deposit the mRNA, also need to be higher. This could already be expected based on the burst-destruction of the intranodal mRNA-loaded microbubbles.

Using the preset “burst” function on the clinical scanner, the microbubble contrast signal is maximally reduced by 50%, indicating an incomplete intranodal microbubble destruction. Only after 6 bursts the mean echo reaches a minimum. On this basis, the transducer that was used in this study (12-5 MHz linear) is likely not ideal for microbubble destruction that could allow effective sonoporation and mRNA transfection. Other clinical transducers that can emit lower frequencies, which are closer to the resonance frequency of the microbubbles and can emit higher powers, could be better suited. In any case, optimization of the various ultrasound parameters such as acoustic pressure, pulse duration and number of pulses, will be required for this purpose [].

Conclusion In conclusion, we were able to show that homemade unloaded as well as mRNA-loaded microbubbles efficiently reach the lymph vessels and nodes upon subcutaneous injection in dogs. Loading of the microbubbles with mRNA-lipoplexes had no significant effect on the distance of microbubble migration from the injection site, nor on the intensity of the observed contrast signals. This shows that theranostic mRNA-loaded microbubbles could have potential for the ultrasound-guided, ultrasound-triggered intranodal delivery of mRNA. However, further research is needed for the optimization of acoustic parameters to most effectively sonoporate target cells of interest. Abbreviations CEUS: contrast-enhanced ultrasound imaging; DC: dendritic cell; SLN: sentinel lymph node; DOTAP: 1,2-dioleoyl-3-trimethylammonium-propane; DOPE: 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine: DSPE: 1,2-distearoyl-sn-glycero-3-phosphoethanolamine; PEG: polyethylene glycol; DPPC: 1,2-dipalmitoyl-sn-glycero-3-phosphocholine; ROI: region of interest; REML: restricted maximum likelihood procedure; CI: confidence interval; MI: mechanical index.

Supplementary Material Acknowledgements Heleen Dewitte is a doctoral fellow of the Institute for the Promotion of Innovation through Science and Technology in Flanders, Belgium (IWT-Vlaanderen). Ine Lentacker and Katrien Vanderperren are postdoctoral fellows of the Research Foundation-Flanders, Belgium (FWO-Vlaanderen). This project was funded through the FWO grant G016513N.

The authors would also like to thank Prof.dr. Jean-Paul Remon for providing the animals, and Prof.dr. Mike Averkiou and Ying Luan for the helpful discussions. Competing Interests The authors have declared that no competing interest exists. References 1. Microbubbles in medical imaging: current applications and future directions. Nat Rev Drug Discov.

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Ultrasound Med Biol. 2012; 38:2174-2185 Author contact Corresponding author: Stefaan C. De Smedt: Laboratory for General Biochemistry and Physical Pharmacy, Ottergemsesteenweg 460, B-9000 Ghent, Belgium. Tel +32 9 264 80 76; Fax +32 9 264 81 89; Email: Stefaan.DeSmedtbe. Received 2014-8-8 Accepted 2014-9-22 Published 2015-1-1.

Background Lymphatic pump techniques (LPT) are used clinically by osteopathic practitioners for the treatment of edema and infection; however, the mechanisms by which LPT enhances lymphatic circulation and provides protection during infection are not understood. Rhythmic compressions on the abdomen during LPT compress the abdominal area, including the gut-associated lymphoid tissues (GALT), which may facilitate the release of leukocytes from these tissues into the lymphatic circulation.

This study is the first to document LPT-induced mobilization of leukocytes from the GALT into the lymphatic circulation. Methods and Results Catheters were inserted into either the thoracic or mesenteric lymph ducts of dogs. To determine if LPT enhanced the release of leukocytes from the mesenteric lymph nodes (MLN) into lymph, the MLN were fluorescently labeled in situ. Lymph was collected during 4 min pre-LPT, 4 min LPT, and 10 min following cessation of LPT.

LPT significantly increased lymph flow and leukocytes in both mesenteric and thoracic duct lymph. LPT had no preferential effect on any specific leukocyte population, since neutrophil, monocyte, CD4+ T cell, CD8+ T cell, IgG+B cell, and IgA+B cell numbers were similarly increased. In addition, LPT significantly increased the mobilization of leukocytes from the MLN into lymph.

Lymph flow and leukocyte counts fell following LPT treatment, indicating that the effects of LPT are transient. Introduction T he lymphatic system collects proteins and excess interstitial fluid into afferent lymphatic vessels. This lymph also carries antigens and antigen-bearing cells from infected tissues to lymph nodes, where antigen-specific immune responses are initiated. The resulting primed lymphocytes then exit the lymph nodes via efferent lymphatic vessels and re-enter the lymphatic circulation.

The thoracic duct is the largest lymphatic vessel, and it transports lymph from most body tissues, excluding the right arm, the right side of the head, neck and chest, and the right lung and lower left lung lobe, which are drained by the right lymphatic duct. Lymph from the thoracic duct enters the blood circulation at the left subclavian vein, allowing primed lymphocytes to enter general blood circulation. This lymphocyte recirculation facilitates interactions of lymphocytes with foreign antigens in blood and tissue, and is an important component of the immune system. Unlike circulating blood, the movement of lymph through lymphatic vessels is not maintained by the pumping of the heart. Instead, lymphatic circulation is maintained through the rhythmic, phasic contraction of lymph vessel walls and external compression of the lymph vessels.

A series of one-way valves along the vessels ensures unidirectional lymph flow toward their junction with the blood circulation., Forces external to the lymph vessels such as respiration, intestinal peristalsis, and muscle contraction facilitate lymph flow. In addition, activities such as exercise,, passive limb movement,, and body-based manipulative medicine techniques,, have been shown to increase thoracic duct lymph flow. Diseases that result in congestion of the lymphatic circulation, such as infection and lymphedema, can inhibit leukocyte recirculation and exacerbate the disease process.

– Therefore, interventions that relieve lymphedema and enhance lymph-tissue recirculation of immune cells, immune products, or pharmaceuticals should aid in the treatment of infectious disease. Specifically, limb elevation and compression garments reduce filtration of fluid from vascular capillaries and accelerate removal of excess interstitial fluid by the lymphatic drainage. This lymphatic drainage can be further stimulated by intermittent, external pneumatic compression. In addition, manual therapies used by osteopathic practitioners, physical therapists, and massage therapists have been reported to reduce lymphedema., Osteopathic physicians believe that removing obstructions to tissue blood and lymph flow is one of the most effective ways to promote and restore health.,, A group of osteopathic manipulations known collectively as the lymphatic pump techniques (LPT) are designed to enhance lymph return from specific areas of the body. These techniques include the thoracic, liver, splenic, and pancreatic pumps, the pedal pump, and the abdominal lymphatic pump., In addition to reducing edema, the increased lymph flow that results from these treatments is thought to accelerate the removal of cellular wastes, toxins, and bacteria from the interstitial fluid.

These lymphatic pump techniques are also reported to enhance immune function.,– In clinical studies, LPT increased vaccine-specific antibodies,, reduced antibiotic use during infection, – and reduced the duration of hospital stay in elderly patients with pneumonia. Collectively, these studies suggest that LPT stimulates immune responses, which may accelerate the clearance of infection. However, the mechanisms by which LPT may enhance immunity and provide protection during infection are still poorly understood. Previously, we demonstrated that abdominal LPT increased leukocyte counts in thoracic lymph; however, the tissue source of these mobilized leukocytes was unknown. Studies to identify the source of thoracic duct lymphocytes indicate that the majority of thoracic duct lymphocytes are derived from the gut-associated lymphoid tissue (GALT).

– Rhythmic compressions of the abdomen during LPT most likely compress regional lymphoid tissues, including the gastrointestinal mucosa, which may release pooled leukocytes into the lymph circulation. The purpose of this study was to determine if, in fact, abdominal LPT increases leukocytes in both thoracic and mesenteric duct lymph, and determine if the mesenteric lymph nodes (MLN) are a source of these leukocytes. Surgical techniques Dogs were anesthetized with sodium pentobarbital (30 mg/kg, i.v.). After endotracheal intubation, the dogs were ventilated with room air supplemented with O 2 to maintain normal arterial blood gases. A femoral arterial catheter provided blood samples to measure arterial blood gases, and this catheter was also connected to a transducer to monitor arterial blood pressure, which remained normal throughout the experimental protocols. A femoral venous catheter was used to administer supplementary anesthetic.

Collection of thoracic duct lymph In six dogs, the chest was opened by a thoracotomy in the left fourth intercostal space. The thoracic duct was isolated from connective tissue and ligated.

Caudal to the ligation, a PE 60 catheter (i.d. 0.76 mm, o.d. 1.22 mm) was inserted into the duct and secured with a ligature. Approximately 60 min following cannulation of the thoracic duct, thoracic lymph was collected at 1 min intervals during 4 min pre-LPT (baseline), during 4 min of LPT, and at 2–5 min intervals for 10 min following cessation of LPT (post-treatment condition). Lymph flow rate was computed from the volume of lymph collected during these time intervals.

Collection of mesenteric duct lymph For collection and analysis of mesenteric lymph, six dogs were surgically prepared for experimentation as described above, except that instead of opening the chest, a midline abdominal incision was made to expose a large mesenteric lymph duct. This duct was isolated, ligated, and a PE60 catheter was inserted into the duct and secured with a ligature. This catheter was exteriorized through the abdominal incision, which was then closed with 2-0 silk suture. Approximately 60 min following cannulation of the mesenteric lymph duct, mesenteric lymph samples were collected, and flow was measured as described above for thoracic duct lymph. Fluorescent labeling of mesenteric lymph nodes in situ Five dogs were surgically prepared for thoracic duct lymph collection as described above, with incisions in both the left chest and in the midline abdomen. After the abdominal cavity was opened, readily visible mesenteric lymph nodes (MLN) were labeled as previously described. Briefly, 12 mg of lyophilized 5(6)-Carboxyfluorescein diacetate N-succinimidyl ester (CFSE) were dissolved in 5 ml dimethyl sulfoxide (Sigma, St Louis, MO).

Then, each MLN was directly injected with 100–200 μl of CFSE, depending on the size of the node. The abdominal cavity was closed with 2-0 silk suture.

The thoracic lymph duct was then catheterized for collection of lymph as described above. Thoracic duct lymph samples were collected at 1, 10, 20, 30, 45, and 60 min after labeling the MLN with CFSE, and these lymph samples were analyzed for CFSE-labeled leukocytes and free CFSE (see ). When the concentration of CFSE labeled leukocytes in thoracic duct lymph had reached a steady state, at 45–60 min post-labeling, lymph was collected during 4 min pre-LPT (min 61, 62, 63, and 64), during 4 min of LPT (min 65, 66, 67, and 68), and for 10 min post-LPT (78 min). The number of CFSE-labeled leukocytes was measured in these lymph samples. Determination of total fluorescence and fluorescing leukocytes in lymph samples Two-color immunofluorescent staining and flow cytometry were performed to identify lymphocyte populations.

Fluorescein isothiocyanate (FITC) labeled anti-canine CD3, phycoerythrin (PE)-anti-canine B cell, PE-anti-canine CD4, AlexaFluor 647-anti-canine CD8, FITC-anti-canine IgA, or FITC-anti-canine IgG monoclonal antibodies (mAb) (Serotech, Raleigh, NC) were used. A total of 10 6 cells were incubated with the mAb as described by the manufacturer.

The cells were washed in staining buffer consisting of Mg 2+-free, Ca 2+-free phosphate buffered saline supplemented with 2% fetal bovine serum (HyClone Laboratories, Logan, UT) and fixed with 0.05% paraformaldehyde until analyzed. Following intranodal labeling of the MLN, lymph was centrifuged and CFSE in supernatants of thoracic duct lymph was measured with a Cary Eclipse spectrofluorometer (Varian Inc., Palo Alto, CA). The excitation wavelength was 480 nm, and the emission signal was collected at wavelengths from 500 to 570 nm. Each sample was scanned three times and a mean value was computed.

Fluorescently labeled lymphoid cells were analyzed using a Cytomics FC 500 flow cytometer (Beckman Coulter, Fullerton, CA). Lymphocyte gates and detector voltages were set using isotype control stained cells, and stained cell populations were seen as distinct peaks or clusters of cells. The proportion of each cell population was expressed as the percentage of the number of stained cells. To determine the total number of a specific lymphocyte population in a milliliter of lymph, their percentage was multiplied by the total number of cells. To determine the flux of a leukocyte population (cells/min), the cell population number was multiplied by the thoracic duct lymph flow.

Statistical analyses Data are presented as arithmetic means ± standard error (SE). Values at individual time points were analyzed and are plotted in figures.

Mean values for pre-LPT, LPT, and post-LPT conditions were computed for each animal; these values were analyzed and are reported in tables. For statistical evaluation, data were subjected to repeated measures analysis of variance followed by a Tukey–Kramer multiple comparisons post test. Analyses were performed with Graphpad Prism version 5.0 for Windows, (GraphPad Software, San Diego, CA).

Differences among mean values with P ≤ 0.05 were considered statistically significant. Abdominal LPT increases leukocytes in thoracic duct lymph During the pre-LPT condition, thoracic duct lymph contained 12.5 ± 2.2 × 10 6 leukocytes/ml, a value consistent with prior measurements in dogs, rats,, and humans. LPT quickly increased leukocyte count in thoracic duct lymph to a peak value 34.0 ± 15.0 × 10 6 cells/ml at 2 min of treatment ( P 0.05). The thoracic duct lymph flow pre-LPT was 0.62 ± 0.12 ml/min, and during the 4 min of LPT, the lymph flow averaged 4.2 ± 0.39 ml/min ( P. Abdominal LPT mobilizes leukocytes into mesenteric duct lymph The average pre-LPT leukocyte count in mesenteric lymph was 6.3 ± 0.67 × 10 6 cells/ml, consistent with values previously reported for sheep, and rats. Thus, basal mesenteric leukocyte count was approximately 50% of that of thoracic duct lymph. LPT quickly increased leukocyte count in mesenteric lymph to a peak value 18.0 ± 4.11 10 6 cells/ml ( P 0.05, vs pre-LPT).

The average pre-LPT, mesenteric lymph flow was 0.35 ± 0.07 ml/min. During the first 2 min of LPT, the duct lymph flow increased to an average of 1.4 ± 0.42 ml/min ( P. Abdominal LPT mobilizes leukocytes from the mesenteric lymph nodes into thoracic duct lymph Immediately following intranodal labeling of the MLN with CFSE, 80%–90% of leukocytes in samples of thoracic duct lymph were labeled with CFSE. This percentage then gradually fell until, by 45–60 min postlabeling, the percentage of labeled leukocytes in thoracic duct lymph was nearly constant at approximately 16%. Thus, in this steady-state baseline condition, about 16% of the thoracic duct leukocytes had originated from labeled MLN.

The initial greater percentage of labeled leukocytes in thoracic duct lymph was most likely due to extranodal labeling of leukocytes with CFSE that had escaped from the injection sites. This view is supported by detection of unbound CFSE in supernatant of samples of thoracic duct lymph collected soon after injection of CFSE. However, by 45–60 min following intranodal injections of CFSE, unbound CFSE was no longer detected in thoracic duct lymph. Thus, by 45–60 min postlabeling, the amount of CFSE in thoracic duct lymph provided a stable index of the contribution of MLN-derived leukocytes to total leukocytes in thoracic duct lymph. This percentage of CFSE-labeled leukocytes in the thoracic duct samples collected were 16 ± 3.1% pre-LPT, 19 ± 4.8% during LPT, and 21 ± 7.7% post-LPT.

If the MLN were not a source of the leukocytes released into lymphatic circulation during LPT, the percentage of CFSE-labeled leukocytes would have decreased markedly during LPT, since thoracic duct flow increased. While LPT did not significantly increase the percentage of CFSE labeled leukocytes in the thoracic duct lymph, LPT did increase the lymphatic flux of CFSE labeled leukocytes (), since thoracic duct lymph flow was increased approximately 4-fold. Considering the flux of leukocytes in the thoracic duct, and the percentage of these leukocytes that were derived from MLN, these data demonstrate that 4 min of LPT produced an incremental mobilization of 27 × 10 6 leukocytes from MLN. Thus, the hypothesis that LPT increases mobilization of leukocytes from the MLN into the lymphatic circulation is supported. Discussion This report provides the first data on the effects of LPT on mesenteric lymph flow and leukocytes in mesenteric lymph. Consistent with prior studies of this laboratory on the effects of LPT on thoracic duct flow and leukocyte counts,,, abdominal LPT produced significant elevations of mesenteric lymph flow and leukocyte counts and flux. While lymphatic leukocyte numbers remained statistically elevated during the entire 4 minutes of LPT, a decline in leukocytes was observed during the last 3-04 minutes of LPT, particularly in the mesenteric lymph.

This was due primarily to reduced lymph flow, and suggests that LPT mobilizes mesenteric lymph from a fluid pool that can deplete within a few minutes of treatment. Mesenteric lymph dynamics are of particular importance since leukocytes from the GALT are transported by the mesenteric lymph circulation.

Processo Penal Esquematizado Pedro Lenza Pdf Download. LPT increased the numbers of IgA and IgG surface-bearing B cells in mesenteric and thoracic duct lymph, demonstrating that LPT is indeed able to mobilize mature, antigen-specific lymphocytes from GALT into the lymphatic circulation. Of the leukocytes in thoracic duct lymph, approximately 16% came from labeled mesenteric lymph nodes.

LPT markedly increased lymph flow with no decrement in the percentage of labeled cells, thus demonstrating the ability of LPT to significantly enhance mobilization of leukocytes from MLN. It is important to note that only 25%–50% of the MLN were labeled; therefore, the actual percentage of thoracic duct leukocytes that originate from the MLN would be much greater than our data indicate. The greatest increase in leukocyte concentrations was observed during the last two minutes of LPT ().

These finding data indicate that it takes approximately 2 minutes for leukocytes released from the MLN to be transported into the thoracic duct. LPT rapidly increased mesenteric leukocyte numbers in lymphatic circulation, suggesting the GALT is a tissue source that is sensitive to LPT. Leukocytes from the spleen exit via the splenic vein and take hours to enter peripheral blood circulation and lymphoid organs. Therefore, it is unlikely that LPT mobilized leukocytes from the spleen into lymphatic circulation within the 4 minutes of treatment. Furthermore, splenectomy does not significantly alter thoracic duct lymphocyte numbers, suggesting that leukocytes from the spleen do not readily migrate into thoracic duct lymph.

Collectively, these findings suggest that spleen is not a source of the leukocytes mobilized into lymph during LPT. While the release of leukocytes into lymph during LPT was transient, 4 minutes of LPT produced a net increase of 6 × 10 8 leukocytes into thoracic duct lymphatic circulation.

Although this increase of leukocytes transported into circulation via lymph during LPT may seem relatively minor compared to the total leukocytes found in blood, increased mobilization of leukocytes could enhance immune surveillance and promote earlier responses to pathogens. Mature lymphocytes bearing antigen-specific receptors recirculate continually from the bloodstream through the peripheral or secondary lymphoid organs, and then return to the bloodstream via the lymphatic vessels., Most adaptive immune responses are initiated when these recirculating lymphocytes recognize specific antigens on the surface of an activated professional antigen presenting cell. If this circulation of lymphocytes is restricted in any way, there could be a delay in the immune response to a pathogen, which could compromise the health of an individual.,,, Therefore, LPT may be an approach to not only to improve lymph flow but, more importantly may also increase the release of lymphocytes into circulation, resulting in earlier and more frequent encounters and responses to a pathogen. The mucous membranes, which line the respiratory, digestive, and urogenital tracts, are a major site of antigen entry., Studies have shown that antigen-specific lymphocytes primed in the gastrointestinal tract can migrate into the lungs and provide protection during pulmonary infection.

– Furthermore, leukocytes from the GALT comprise a different array of immune specificities and responses than other tissues due to the antigens that they encounter. Thus, LPT may not only enhance circulation of leukocytes, but it may also facilitate the trafficking of local immune responses to other tissues, such as the lungs, that may benefit from their presence during infection. In conclusion, clinical reports suggest that LPT, can stimulate immune responses, – which may accelerate the clearance of infection. – In support of this notion, we have demonstrated that LPT increases the release of leukocytes from the GALT into lymphatic circulation.

The information gained from this study provides a rationale for the use of LPT to enhance immunity.